Chemicals
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2-Dimethylaminoethanol (DMAE)
C050 - 2-Dimethylaminoethanol (DMAE) FLAMMABLE, CORROSIVE Causes burns Harmful by inhalation, in contact with skin and if swallowed. May cause sensitisation by skin contact. Lachrymator In case of contact with eyes, rinse immediately with plenty of water and seek medical advice. Immediately take off all contaminated clothing. Wear suitable protective clothing, gloves and eye/face protection. SPILL OR LEAK PROCEDURES Absorb on sand or vermiculite and place in closed containers for disposal. Waste disposal method Dissolve or mix the material with a combustible solvent and burn in a chemical incinerator equipment with an afterburner and scrubber.
Agarose I
EMS10205, EMS10207 TIPS FOR PREPARING When agarose is placed in a buffer such as TAE (Tris/Acetate/EDTA) or TBE (Tris/Borate/EDTA), it is generally insoluble. However, when this agarose solution is heated, the agarose particles become hydrated and thus go into solution. This hydration process is time-dependent, and different types of agarose will have varying hydration points. EMS’s agaroses are extremely pure and comprised of ultra-fine particles. This ultra-fine structure, EMS agaroses will have a faster rate of hydration than other type of agaroses. End-users who have used other brands of agarose in the past may mistakenly boil EMS agaroses much longer than is needed, which results in a thick gelatinous solution that is difficult to cast and brittle when polymerised. Guide for Preparing EMS Agaroses Preparation of a typical 1% agarose gel 1X TBE buffer. In an appropriate container (an Erlenmeyer flask at 2-4x the volume of the desired gel volume is optimal), slowly add agarose crystals to your buffer solution while gently swirling. This will help to eliminate clumping of the agarose. Record the weight of the flask containing the buffer and agarose. Heat the solution in a microwave on high power for 30 seconds (for smaller or larger volumes, increase or decrease heating times proportionally to volume size). Heating times will vary depending on your microwave oven (wattage), size of the flask used and the % agarose. Swirl the agarose solution gently to re-suspend the particles. Heat the solution another 30 seconds on high power, remove and swirl the agarose solution. Place the solution back in the microwave and heat on high power until the solution just starts to boil (boiling point will probably take 10-35 seconds). Use caution when handling the hot flask. Microwave solutions may become superheated and can be boil vigorously when moved or touched. After removing the boiling solution from the microwave oven, allow to cool briefly (1-2 minutes) at room temperature, then gently swirls the solution to release entrapped air (some air bubbles will remain). Place the agarose solution back in the microwave, heat on high power and let the solution boil for approximately 15 seconds. Inspect the solution for agarose crystals (they will appear as floating ‘lenses’) while gently swirling. If there are particles present, repeat this step until all crystals are dissolved. Once the agarose is completely in solution, again weight the flask to check for water loss by evaporation. Replenish with water as necessary (until the weight of the flask and its contents equal the original weight). Gently swirls the solution. In general, it is advisable to allow any agarose solution to cool to ~ 50-55°C on the lab bench prior to pouring into a prepared apparatus. This is conductive to a more uniform pore size and will prevent the warping of your gel apparatus. Before pouring the gel, gently swirls the agarose solution to help dissipate most of the remaining air bubbles. Pour the gel into the prepared casting unit. Usually, horizontal gels should be 3-5mm thick. Immediately after pouring, check to see that there are no air bubbles under or between the teeth of the gel comb. Allow the gel to completely polymerise at room temperature (about 30-45 minutes) before running your samples. For further information on agarose gel electrophoresis, see Sambrook et al. (Chapter 6). If you are preparing a different type of agarose (i.e., a higher concentration or a different volume), the most important things to remember are: Gently swirls your agarose solution at least twice before you bring the solution to a boil. Once the boiling point has been reached, observe your solution after each 10-15 seconds, boiling intervals very closely (depending on concentration and volume) SEPARATION OF BIOLOGICAL MOLECULES BY GEL ELECTROPHORESIS Protocol Overview In a flask 2-4 times greater than gel volume add agarose to buffer with constant swirling. Weigh flask Microwave on high for 30 seconds Swirl solution Microwave on high for 30 seconds Swirl solution Microwave on high until solution boils Remove and allow to cool for 1-2 minutes before swirling solution again Boil again for 15 seconds Check for crystals and repeat boiling until solution is homogenous Weigh flask and add water to return to original weight Cool to 50-55°C Swirl and pour into casting stand Allow 30-45 minutes for gel to solidify Microwave instructions: Recommended for agarose concentrations ≤ 3%. Agarose must be uniformly dispersed in buffer prior to hydration to avoid clumping. Agarose I tablets should be crushed into a powder with a clean rod or spatula prior to dispersion. Determine gel volume and agarose concentration: Estimate volume: Volume = surface area of the casting chamber X gel depth Optimal resolution is usually obtained on gels 3-4 mm thick. In an appropriate container (an Erlenmeyer flask at 2-4 times the volume of the desired gel volume) slowly add agarose crystals to your buffer solution while gently swirling the flask. Weigh the flask containing the buffer and agarose. Heat the solution in a microwave on high power for 30 seconds. (For smaller or larger volumes, increase or decrease heating times proportionally to volume size). Heating times will vary depending on the wattage of your microwave oven, size of the flask used and the agarose concentration. Swirl the agarose solution gently to re-suspend the particles. Heat the solution another 30 seconds on high power, remove and swirl the agarose solution. Place the solution back in the microwave and heat on high power until the solution just starts to boil (boiling point will probably take 10-35 seconds). Caution: Handle the hot flask very carefully. Microwaved solutions may become superheated and boil over when moved or touched. Remove the boiling solution from the microwave oven, allow to cool briefly (1-2 minutes) at room temperature. Gently swirl the solution to release entrapped air (some air bubbles will remain). Place the agarose solution back in the microwave, heat on high power and let the solution boil for approximately 15 seconds. Inspect the solution for agarose crystals (they will appear as floating “lenses”) while gently swirling. If there are particles present, repeat this step until all crystals are dissolved and the solution is transparent. Once the agarose is completely in solution, reweigh the flask to check for water loss by evaporation. Replenish with hot distilled, deionised water until the weight of the flask and its contents equal the original weight. Gently swirl the solution. Allow the agarose solution to cool at room temperature to ~50-55°C before pouring the gel into the prepared casting stand. This will result in a gel with a more uniform pore size and prevent warping of the gel apparatus. Notes: Preparation of any agarose solution in the microwave requires constant attention to prevent the solution from boiling over. Gently swirl agarose solution at least twice before bringing the solution to a boil. Once the boiling point has been reached, observe the solution after each 10-15 second boiling interval until particles are no longer visible. Hotplate/magnetic stirrer instructions: Particularly recommended for agarose concentrations between 4%-5%. It may be used for lower concentrations as well. Follow Step 1 for microwave instructions (above). With heat off, place an appropriate container (an Erlenmeyer flask at 2-4 times the volume of the desired gel volume) containing buffer and a PTFE coated stir bar on the magnetic stirrer. With rapid stirring to prevent formation of clumps slowly add agarose to your buffer solution. Weigh the flask containing the buffer, agarose and stir bar. With rapid stirring bring solution to a boil. Maintain a gentle boil until agarose is completely dissolved, about 3-10 minutes. Inspect solution for agarose crystals and continue boiling if necessary. Follow steps 9-13 in microwave instructions above (above). Autoclave Instructions: Particularly recommended for agarose concentrations between >5%. It may be used for lower concentrations as well. Caution: Agarose solutions containing Ethidium Bromide or other mutagenic intercalating stains should not be autoclaved to avoid apparatus contamination. Follow steps 1-6 for microwave instructions (above). Cover flask with aluminium foil and autoclave at 121°C for 15 minutes. Remove from autoclave and allow to cool briefly. Weigh the flask and add warm distilled water to return to original weight. Allow to cool to 50-55°C before pouring into casting stand. Gel Casting Assemble casting stand according to the manufacturer’s instructions. • Casting stand should be level. • Comb teeth should be examined for dried agarose and cleaned with hot water and rinsed in distilled, deionised water prior to use. When agarose solution cools to ~55°C, gently swirl to help dissipate most of the remaining air bubbles. Pour the gel into the prepared casting unit to a depth of 3-4mm. Immediately after pouring, insert comb and check to see that there are no air bubbles under or between the teeth of the gel comb. Allow the gel to completely solidify at room temperature (about 30-45 minutes). Note: To achieve optimal resolution with Agarose SFR™: Allow SFR™ agarose gels to solidify completely at room temperature. The gel should then be placed at 4°C for 30 minutes prior to loading and running.
Alconox Detergent
M030 Mixing Directions for Alconox Detergents Dilute detergent (see chart below) using warm water (about 50°C) or hot (about 60°C) water. Ambient temperature water may be used, especially for presoaking. For difficult soils, use very hot water (above 65°C) and use double the recommended amount of detergent. If you are to reuse the cleaning solution, make up fresh solutions as frequently as needed. Soaking Recommended Products: Alconox, Liqui-Nox, Citranox, Terg-A-Zyme, Alcojet, Det-O-Jet, Luminox, Detergent 8, and Citrajet. Typical Use: To clean small items – hospital catheters and tubes, small metal parts and large tank interiors, including pharmaceutical and other blending tanks. An excellent pre-treatment method for loosening soils and preventing drying – especially for labware or medical instrumentals prior to further cleaning Advantages: Little physical effort or expense. Concerns: Extremely dirty articles or difficult soils may require further cleaning. Directions: Soak, completely submerged in solution until clean (may take several hours depending on soil type) Remove and rinse thoroughly (see Rinsing) Product Form Dilution (%) Recommended Mixture Temp Usual Wash Temp Protective Gloves Eye Protection Alconox Powder 1 10g/L Ambient Warm Desired Desired Terg-A-Zyme Powder 1 10g/L Ambient Max 54°C Desired Desired Liqui-Nox Liquid 1 10ml/L Ambient Warm Desired Desired Citranox Liquid 1-2 10-20ml/L Ambient Hot Required Required Detergent 8 Liquid 2-5 20-50ml/L Ambient Hot Required Desired Luminox Liquid 2-5 0-50ml/L Ambient Hot Required Desired Alcojet Powder 1 5-10g/L Warm Hot Required Desired Det-O-Jet Liquid 1 5-10ml/L Ambient Hot Required Required Alcotabs Tablet - (1 tablet per use) Ambient Ambient N/A N/A Citrajet Liquid 1-2 10-20ml/L Ambient Hot Required Required MANUAL CLEANING Recommended Products: Alconox, Liqui-Nox, Citranox, Terg-a-Zyme, Alcojet, Det-o-Jet, Luminox, Detergent 8, and Citrajet. Typical Use: For cleaning small articles such as medical examination instruments, labware or circuit boards, and large articles such as process equipment. Advantages: Versatile, inexpensive, effective. Concerns: Time consuming and labor-intensive. May not be effective on difficult-to-reach areas requiring pre-soak, ultrasonic, or machine cleaning. Directions: Make up cleaning solution as in mixing directions, or use undiluted detergent on a warm, wet cloth or sponge for non-abrasive scouring. Clean as follows: wet the article with solution by dunking or using a soaked cloth or sponge Clean with a cloth, sponge, cotton swab, brush or pad that agitates surface soils without marring the surface. Rinse thoroughly (see Rinsing). Wear gloves, eye protection, and other safety equipment if recommended. ULTRASONIC CLEANING Recommended Products: Alconox, Liqui-Nox, Citranox, Terg-a-Zyme, Alcojet, Det-o-Jet, Luminox, Detergent 8, and Citrajet. Typical Use: To clean large batches of articles or for fast, convenient cleaning. Advantages: Fast, effective, penetrating cleaning. Concerns: Capital cost, material tolerance for ultrasonic agitation. Directions: Make up detergent solution in a separate container. Add cleaning solution, run machine for several minutes to de-gas solution and allow heater to come up to temperature. Place groups of small articles in racks or baskets. Align irregularly shaped articles so the long axis of any part faces the ultrasonic transducer (usually the bottom). Immerse articles to be cleaned for 2-10 minutes, or longer, as needed. Remove and rinse thoroughly (see Rinsing). CLEAN-IN-PLACE Recommended Products: Alconox, Liqui-Nox, Citranox, Terg-a-Zyme, Alcojet, Det-o-Jet, Luminox, Detergent 8, and Citrajet. Typical Use: For pipe, tank, and filtration systems. Advantages: Assures clean systems without disassembly. Concerns: Good circulation in system. Directions: Make up cleaning solution as in mixing directions. Circulate solution slowly for at least 30 minutes. Allow several hours for large systems (thousands of gallons), especially with ambient temperature water. Drain by pumping in one full system capacity of water. Rinse by circulating and draining at least two times the system's water capacity. Some filtration units may require more rinsing. MACHINE WASHERS Typical Use: For high-volume cleaning using washer-sanitisers, ware-washers, conveyor-washers, or spray and pressure washers. Advantages: Fast, effective, high-volume cleaning. Directions: Load articles into racks so that open ends face toward spray nozzles. Place difficult-to-clean articles with narrow necks and openings near the center of the rack, open-side down, preferably on special racks with spray nozzles pointing directly into them. Minimise touching between articles. Group small articles in baskets to prevent dislodging by spray action. Use only low-foaming detergent as per machine manufacturer dose instructions. If no instructions, use a 1% solution or 10g/L of wash water. Use more or less as needed. Use hot water (above 60°C). Most machines have at least three rinse cycles (see Rinsing). Refer to machine manufacturer's directions. AUTOMATIC SYPHON PIPETTE WASHING Recommended Products: Alcotabs. Typical Use: Washing pipettes in laboratories. Advantages: Effective batch pipette cleaning. Concerns: Pre-soak pipettes for best results. Directions: Completely immerse pipettes immediately after use in a pre-soak solution. When ready to clean, drop an Alcotab into bottom of washer. Place pipettes in holder into the washer. Turn on cold or warm water at a rate that will fill the washer and completely cover all pipettes, then drain to the bottom during each cycle. Run water until Alcotab has completely dissolved. Continue running water to rinse thoroughly (may take an hour to complete washing and rinsing). For analytical or tissue culture work, use distilled or deionised water for final rinse. Rinsing Don't neglect the rinse! Use ambient, warm, or hot water. A running water rinse directly contacting all surfaces for at least 10 seconds on each surface is recommended. If not practical, use a series of three or more agitated soak rinse tanks or at least two counter-flow cascade rinse tanks. For large surfaces, several passes with a clean cloth or sponge soaked with rinse water followed by a clean, dry, absorbent wipe can work. In machine cleaning, after washing, there should be at least three rinse cycles. Tap water is suitable for many rinsing applications. Give medical and surgical instruments a final rinse in distilled or deionised water. In laboratories, rinse tissue culture and analytical ware with deionised or distilled water. Rinse trace organic analytical ware in distilled or organic-free water. Give trace metal or inorganic analytical ware a final rinse with deionised water. Rinse pharmaceutical equipment according to good manufacturing practice with whichever is required: potable, deionised, distilled, sterile, pyrogen-free or injectable water. Rinse electronic circuit boards and non-conducting electronic devices with deionised water. Sensitive optical or precision manufactured parts may require final rinses in deionised or distilled water. Food processing equipment must be rinsed with potable water. Drying Drying can affect residues and corrosion. Impurities from rinse water can be deposited during evaporation. To minimise this, dry with techniques that physically remove rinse water from the substrate such as absorbent wiping, forced air, or air knives, azeotropic solvent drying such as isopropyl alcohol final rinse and dry, or vacuum drying that may also evaporate residues. Water, and particularly high-purity rinse water can be corrosive to metal substrates during heating and air-drying. The use of physical removal drying techniques or the addition of corrosive inhibitors (with the tolerance of corrosion inhibitor residues) to the rinse water can help minimise corrosion. Bath life monitoring, extension and control For the highest levels of critical cleaning only freshly made up solutions should be used for cleaning to avoid any potential for cross contamination. For industrial critical cleaning applications high levels of cleaning can still be achieved with extended bath life. In general, a pH change of 1 pH unit towards neutral indicates an exhausted cleaning solution. Bath life can be extended by physical filtration of particulates and cooling and settling of sludge and skimming of oils. Bath life can be extended by adding one half as much detergent of the initial load after partially depleting the cleaning life of a bath. Under frequent daily use, detergent solutions can rarely be used more than a week even with these bath life extension techniques. Conductivity, pH and, % solids by refractometer can be used to control bath detergent concentration. Free alkalinity titration can also be used to control bath life of alkaline cleaners where the soil being cleaned depletes free alkalinity – as is often the case with oily soils. Titrate a fresh solution to determine new solution free alkalinity. Titrate your used solution to determine the percent drop in free alkalinity. Add more detergent to the bath to bring the free alkalinity back to the new solution free alkalinity. For example if your initial solution is made up with 100 ml of cleaner concentrate and you observe a 25% drop in free alkalinity, you should try adding 25 ml of cleaner concentrate to recharge your solution. You should perform a new free alkalinity titration to confirm your recharge the first few times you use this recharging method to be sure that the detergent you are using is linear with respect to free alkalinity depletion. This form of bath life extension cannot run indefinitely, sludge will eventually form. Fresh solutions must be periodically made up. Corrosion Inhibition Corrosion during cleaning is accelerated by the same things that accelerate cleaning: heat, aggressive chemicals, time, and agitation. In approximate order of importance, to reduce metal corrosion concerns, use less heat, lower pH detergents, shorter cleaning time, and less agitation. In general use the mildest pH detergent to limit metal corrosion. Higher pH detergents such as Alcojet have special corrosion inhibitors that often allow it to be used successfully to clean soft metals such as aluminum. In approximate order of importance, in general to reduce plastic corrosion, use less aggressive cleaners that have less solvent or surfactant character or use lower concentrations of those cleaners, use lower cleaning temperatures, use less contact time, and finally use less agitation. With aqueous cleaning metal corrosion can occur during rinsing and drying. Corrosion inhibitors can be added to rinse water provided that any associated inhibitor residue does not interfere with the surface being cleaned. Keeping the surfaces cleaned hot with hot rinse water and using rapid heat or vacuum drying can accelerate drying and minimise metal corrosion. Forced air drying and air knives that physically remove rinse water can minimise drying corrosion. Drying with hot oxygen-free gas such as nitrogen can also control corrosion during drying. With mild steel you can have "flash rusting" when you rinse with hot water and dry with hot air. In some instances, by lowering the water temperature or drying temperature, corrosion can be avoided on mild steel. For instance in a case where flash rusting on mild steel had been occurring using 66°C rinse water and ambient air drying, it was found that flash rusting could be avoided by using 49°C rinse water in place of the 66°C rinse water. Cleaning measurement procedures In any industrial or processing application, "clean" means clean enough to avoid later problems. What happens if the surface is not clean? You can choose the best testing methods for measuring cleanliness and assuring quality of product: Water-break: (10s of mg/cm2 level) Use running water and let it sheet across the surface. Observe if any breaks in the water occur due to hydrophobic (water-fearing) residues. The water break test is a fairly crude test that is suitable for detecting films of process oils and heavy fingerprints. It will not readily detect non-hydrophobic residues. This test is often used for parts washing; it may not always be suitable for precision cleaning applications. Atomiser: (10s of mg/cm2 level) A variation of the water-break test, this requires observing whether a gently sprayed water mist deposits uniformly or whether water repulsion occurs (usually due to a hydrophobic soil). The atomiser test is slightly more sensitive to hydrophobic soils than the water break test. In the water break test, the kinetic energy of the flowing water may overcome a hydrophobic residue, where in the atomiser test, you may be able to see the results of a little droplet of water being repelled by a hydrophobic contaminant. Oil-soluble fluorescence: In this test, the cleaned part is dipped into a fluorescent oil. If oily residues are present, they will be extracted into the florescent oil that can be easily observed under ultraviolet light. Of course, using oil-soluble fluorescence requires cleaning the part again after each test. Oil-soluble florescence is a highly sensitive method and can give very dramatic visible results of oil residue. The problem with this test is that it really only shows how well your process removed oily soils, it does not reveal particulate or inorganic soils. Non-volatile residue: (10s of mg/cm2 level) extract contaminants from a part using a volatile solvent and then evaporate the solvent in a pre-weighed container and then weigh again after evaporation to detect the weight of volatile residues. Extraction: (mg/cm2 level) A particularly excellent method for detecting detergent residues. Use a solvent-soaked glass filter paper or polyester swab to wipe surface. Extract or digest filter paper. Use trace analysis on the extract. Can be quantitative if you wipe a known area. Extraction method can be highly sensitive to a wide range of possible soils and residues. The limitations of this method are that an appropriate solvent has to be used to extract whatever type of soil might be there. If you were doing trace analysis for detergent residues you would use water as your solvent. The sensitivity of an extraction test depends on your method of trace analysis. The types of trace analysis often used are UV visible spectrophotometry, total organic carbon (TOC) analysis, high performance liquid chromatography (HPLC), atomic absorption (AA) or inorganic residues, and liquid chromatography (LC) and filtration with microscopic filter examination. Oil evaporation: (10s of mg/cm2) For filmy residues, a few drops of organic solvent can be deposited on the surface and then removed via pipette and placed on a watch glass. If any filmy residues are present, you should observe a characteristic ring of organic-material deposits. Gravimetric: (mg/cm2 level) With small parts of known weight, the amount of excess weight indicated the amount of soil present. Surface-energy: (mg/cm2 level) Any hard, flat material has a characteristic surface energy. As a result, a deposit of a known volume of pure liquid (typically deionised water) will form a droplet of predictable size based on the amount of that energy. Measuring droplet size will determine surface cleanliness. Generally, hydrophobic soils create smaller droplets; hydrophilic soils, larger ones. The surface-energy test is far more sensitive than the atomiser or water-break test and has the advantage of being able to detect both hydrophilic and hydrophobic soils. The problem with the surface energy test is it only tests the surface directly underneath the droplet where you are measuring. If you have a broad, large surface where measuring one little area of it would be representative of the level of the cleaning that went on the entire surface, this can be an excellent and highly sensitive method. If you have a part that has lots of cracks and crevices and holes, the surfaces that are accessible for you to place a droplet of liquid to measure surface energy may not be representative of the types of soils that may be hidden in any cracks, crevices, or blind holes. Contact-angle: A variation of surface energy testing is contact-angle, defined simply as the relationship of certain forces when a liquid stabilises on a part's surface. This method can be used to determine cleanliness, since the properties of various contaminated surfaces are reflected by different contact angles. Contact-angle measurement is especially suited to manufacturing operations, such as wire bonding on PCBs or the application of thin films on quartz glass which require a cleanliness test that is nondestructive. Vapors from the process itself, such as vacuum or diffusion pump oils, various process chemicals, or even human perspiration, are all contaminants whose presence can be detected by changes in the contact angle. Accepted methods of measuring contact angles to determine surface characteristics include the inverted bubble, Wilhelmy Plate, and sessile drop techniques. The latter is the most widely practiced quality control technique, as it is relatively quick and requires minimal investment in time and financial resources. Several "real world" examples will help illustrate what can be learned from the use of the contact angle measuring technique to determine the existence of impurities on a surface. LCD (Liquid Crystal Display) panel surfaces contaminated with organic matter will be less accepting of a variety of films, such as metals and protective layers, resulting in poor manufacturing yields. Sources of such contaminants include the vapor of process materials, chemicals, and human perspiration. Very thin organic contaminants several monolayers in thickness (greater than 10 angstroms) can be evaluated using the contact angle technique. In fact, it is generally agreed that the wetting behavior involves only the last layer or two atoms on either side of a solid's interface. The water contact angle correlates the "cleanliness" of the surface to the adhesion of the copper deposited onto the surface of the LCD. Water contact angles can be used in various processes to determine contamination levels, predict cleanliness and adhesive bond strengths, and monitor cleaning operations. Whether you are checking the moisture effects on silicon wafers or LCD quartz panel glass metal adhesion, all that is needed is an understanding of the basic theory involved and proper measurement techniques. As with other cleanliness verification techniques, special tips for contact angle measurement, if practiced, will increase the repeatability and validity of the measurement results. The following tips were helpfully compiled: Use gloves when handling the samples to be measured. Organics, such as finger oils, cosmetics, and other contaminants, will skew the contact angle results. Note the nature of the droplet after applying it to the surface. Wait until the droplet has ceased its advancement and no more change in lateral movement has occurred. Measure this time interval, making sure that you wait this period of time after every measurement. Retain consistent time intervals between the placement of the droplet and its measurement. Use medical-grade, ultra-purified deionised water from a laboratory supply house in order to have a consistent measuring liquid. This will limit the number of measurement variables. Use test liquids of larger surface tension than the solid's surface energy in order to obtain easy-to-read results. Neutralise the effects of static charges on substrates. Substrates that are electrostatically charged can skew contact angle readings up to 5 degrees. Accurately control liquid droplets so that they are repeatedly deposited onto the sample. Gently move the sample to the liquid droplet formed at the end of the syringe/dispenser in order to minimise gravitational effects. In the case of very high contact angles, you may find it difficult to adhere the droplet from the needle to the solid sample. Use a PTFE-coated needle of a higher gauge (smaller inner diameter). In case of very low contact angles, use the highest possible needle gauge (smallest inner diameter needle) for controlling very small droplet volumes onto the sample. Other methods, such as visual examination using a stereomicroscope, fiber optic light, or black light, are also useful in determining the existence of cleaning process residues. For extreme low level residue detection at the many ug/cm2 level, surface analysis techniques such as Auger electron spectroscopy (AES), electron spectroscopy for chemical analysis (ESCA), Fourier transform infrared (FTIR), secondary ion mass.
Antibody Diluent
EMS62713, 62714, 62715 Instructions All antibody preparations have some potential to produce a non-specific reaction in the assay; it originates from: Non-specific antibodies that are present in some proportion in any polyclonal antibody preparation, including ones that are affinity-purified Low specificity antibodies among specific ones in polyclonal Fragments of fallen apart IgGs in stored preparations, including monoclonal Separate heavy and light chains of specific antibodies, which are produced by most hybridomas All these are capable of non-specifically binding to molecules on tissue sections, blots, fixed cells and other objects for immune detection. In the case of retrieved formalin sections, the risk of non-specific reaction is increased, since the proteins comprising the tissue sections are denatured during HIER. This makes many domains accessible that are charged. They are also capable of binding the test immunoglobulins in a non-specific manner. The standard means of blocking non-specific binding of specific antibody preparation is as follows: Add irrelevant protein, other serum, casein, etc. However, in most cases, many who have tried this found that increasing concentration of such blocking agent leads to a great reduction of specific reaction, as well. This is a result of the lack of blocking molecules binding to access sites on section and thus sterically blocking the access to specific antibodies to epitopes of interest. All of our buffers at Electron Microscopy Sciences are developed for immune assays and contain short (0.6-2 kD) peptides that are capable of block effectively to non-specific reactions. These do not affect the specific binding of antibody. Properties Antibody Diluent used in the dissolving of primary and secondary enzymes and antibodies to required working concentrations. Presentation Antibody Diluent is supplied as a ready to use solution in the following volumes: 50, 125 or 500 ml. The solution has a clear colourless appearance. Application Electron Microscopy Sciences recommends this product for research and diagnostic pathology, especially for HIER retrieved sections and polyclonal antibodies. Use in IHC Use to dilute primary polyclonal or monoclonal antibody to reduce non-specific binding of the primary antibody. The diluent can also be used to dilute secondary, enzyme labelled or fluorochrome-labelled secondary antibody. In the case of overdiluting antibodies to reduce the background staining, consider the use of a higher concentration (2-4 times) when using Antibody Diluent, in order to improve the sensitivity or antigen detection. Use in Other Applications Antibody Diluent can also be used to prevent non-specific binding of reagents and to improve sensitivity in the following: Immuno-PRC Western Blotting Protein Arrays Immunofluorescent staining of tissue sections and fixed cells Flow cytometry on fixed and permeabilised cells Use as antibody diluent. For Immuno-PCR and protein arrays, the diluent can also be used as a washing buffer to rinse the reagents between immunostaining stages. Stability and Storage Stable for 2 years when stored unopened at +4°C. Every lot is issued with a certificate indicating the expiration date. After opening, however, store at +4°C in the refrigerator and be sure to use within 6 months.
Antibody Diluent, Frozen Sections
EMS62713-01, 62714-01, 62715-01 General Staining Protocol for Frozen Sections Please read the entire procedure before starting the staining procedure. Perform all incubations in a humid chamber and do not allow sections to dry out. Isotype and system controls should also be run and must be matched to the isotype of each primary antibody to be tested. Materials Needed Phosphate Buffered Saline (PBS) H2O2 Solution Antibody Diluent for IHC Streptravidin-Horseradish Peroxidase DAB Substrate Kit Hematoxylin Bluing Reagent Graded Alcohols Xylene For your convenience, our Ig HRP detection kits can be used to perform the immunohistochemical staining. The Anti-Hamster Ig, Anti-Mouse Ig, and Anti-Rat Ig HRP detection kits are also available, and include Biotinylated Secondary Antibody, Antibody Diluent Buffer, Streptavidin-HRP, DAB Buffer, and DAB Chromogen. Instructions Label slides with a solvent-resistant pen and demarcate the tissue if required. Rinse slides three (3) times in PBS to remove the tissue-freezing matrix. Block endogenous peroxidase activity by incubating the slides in 0.3% H2O2 solution in PBS for 10 minutes. Rinse slides three (3) times in PBS, 2 minutes each time. Block non-specific binding by incubating with blocking buffer (10% serum from host species of secondary antibody diluted in PBS or 10% FBS in PBS) for 30-60 minutes at RT in a humidified chamber. Dilute the primary antibody in the Antibody diluent for IHC. Alternatively, a buffered solution with a source of protein can be used as antibody diluent. Apply the diluted antibody to the tissue sections on the slide. Incubate for 1 hour at RT in a humidified chamber. Rinse slides three (3) times in PBS, 2 minutes each time. Dilute the biotinylated secondary antibody in the Antibody diluent for IHC. Alternatively, a buffered solution with a source of protein can be used as antibody diluent. Apply to the tissue sections on the slide and incubate for 30 minutes at RT. Rinse slides three (3) times in PBS, 2 minutes each time. Apply the Streptravidin-Horseradish Peroxidase pre-diluted to the tissue sections on the slide and incubate for 30 minutes at RT. Rinse slides three (3) times in PBS, 2 times each. Prepare DAB substrate solution by adding 1 drop of DAB chromagen to every 1 ml of DAB buffer. WARNING: DAB is a suspect carcinogen. Handle with care. Wear gloves, a lab coat, and eye protection. Drain PBS from slides and apply the DAB substrate solution. Allow slides to incubate for five (5) minutes or until the desired colour intensity is reached. Wash three (3) times in water, 2 minutes each time. Counterstain slides: Dip twice in Haematoxylin Rinse thoroughly in water Dip twice in Bluing reagent or dilute ammonia water Rinse thoroughly in water Dehydrate through 4 changes of alcohol (95%, 95%, 100%, and 100%). Clear in 3 changes of xylene (or xylene substitute) and coverslip using mounting solution.
Aqua-Hold Pap Pen
ID310 Intended use: The Aqua-Hold pap pen is a special marking pen that provides a thin film-like barrier when a circle is drawn around the specimen. This barrier creates the proper surface tension to hold antibody solution within the target area on the slide. Aqua-Hold is very effective for Immunostaining procedures by the Peroxidase-Antiperoxidase (PAP) method, ABC method, B-SA method, immunofluorescence method, ASD method, Enzyme method and Frozen Section method. The surface tension provided by the barrier ensures that only a small amount of antibody solution is needed. Two specimens, separated by two circles, can be applied to the same slide. The pen contains a special formulation that is water repellent and stable for long time period. It can be removed, if desired, by xylene after the staining procedure is completed. Instructions for use: To start use of pen, take off cover and press felt tip down on a glass slide several times on a flat surface until a flow of material is seen. It might be heavy in flow. If so, wait until the felt tip is fully saturated with film material (turns a darker brown) and wipe off the excess with paper towel. The Aqua-Hold is now primed and can be used to draw the desired barrier on the slide. For Paraffin sections: Deparaffinise and rehydrate the section according to accepted procedures. For proper adhesion of the barrier circle, the glass slide surface should be clean and dry. Remove the slide from water or buffer solution; dry around tissue section with water absorbent paper; encircle the section with the pen. Allow the circle to dry for approximately 1 minute at room temperature and return the slide to water or buffer. If trypsinisation or another proteolytic digestion is to be used, apply the film barrier after that step is completed. For Frozen Section: The film circle may be applied on the glass surface (at room temperature) either before fixation or prior to the first immersion in the water or buffer. If the slide is treated with hydrophile, soak the slide in 0.1N hydrochloric acid for 30 seconds. Wash it with water and dry. This will help prevent the film guard from lifting. Cautions: The solvent is flammable and possibly toxic. Use the pen in a well ventilated area. The Aqua-Hold pen may need further priming after sitting without use. Refer to instructions for use above to continue using the pen.
Araldite 502
Araldite 502 is an epoxy resin used as an embedding medium for electron microscopy. It yields a light gold colour, hard block. Tissues to be embedded in Araldite 502 can be dehydrated with ethanol or acetone. These solvents must be pure and after the third absolute ethanol/acetone change, prior to mixing with liquid resin, a transitional solvent Propylene Oxide (PO), was originally recommended. PO is expensive and is a recognised carcinogen. If you wish to avoid using PO, then make an additional change using pure ethanol or acetone which has be stored with dried Molecular Sieve 4A in a well-sealed jar. Use roughly 10% by volume, then the molecular sieve will further reduce water within organic solvents and the resulting blocks will not be soft or sticky due to water contamination. Tissues to be embedded in Araldite 502 can be dehydrated with most commonly used organic solvents. However the application of a transitional solvent, such as Propylene Oxide, is advisable because epoxy resins are more soluble in propylene oxide. RECOMMENDED PROCEDURE Fixation Tissues can be fixed in a wide range of fixatives. One of the more commonly used fixatives is an Aldehyde (i.e.: Glutaraldehyde) followed by Osmium Tetroxide. Dehydration There are many different dehydration schedules that can be followed. A typical one is as follows: 70% Ethanol for 5 minutes 95% Ethanol for 5 minutes 100% Ethanol for 5 minutes 100% Ethanol for 15 minutes 100% Propylene Oxide for 5 minutes 100% Propylene Oxide for 15 minutes Note: Longer times may be required for some samples. Times are fairly arbitrary, but compounding short-cuts may cause problems. Sensitive specimen (fresh water algae) would be processed using very weak buffers and starting dehydration no higher than at 30% and applying more, smaller increases in solvent concentration. Ingredients Araldite 502 - 20ml Dodecenyl Succinic Anhydride (DDSA) - 22ml Benzyldimethylamine (BDMA) - 1-1.2ml BDMA is recommended for better penetration and stability, however you could also use 2,2-Dimethoxypropane (DMP) - 0.63-0.84ml For larger batches simply increase each component proportionally. ProSciTech Pty Ltd offers an Araldite Embedding Kit as a cheaper alternative to buying each of these chemicals separately. Mixing Instructions Slight variations of the accelerator (DMP-30 or BDMA) will drastically affect the colour and brittleness of the block. Prior to measuring and mixing, the resin and the anhydride should be warmed (60°C) to reduce their viscosity. Thorough mixing is imperative to be able to achieve uniform blocks. The final block can be made harder by replacing some of the DDSA with Methyl-5-norbornene-2,3-dicarboxylic anhydride (NMA) (0.5mL of NMA for each 1.0mL of DDSA). Although the mixture can be stored for up to 6 months at 4°C, it is highly recommended that freshly prepared embedding medium always be used. If you choose to store the mixture, you should warm it thoroughly prior to adding the accelerator. Infiltration It is recommended that for all of the infiltration steps a specimen rotator be used. Drain the tissue of most of the propylene oxide, leaving a little so the tissue does not dry out. Replace the solvent with a 1:1 solution of propylene oxide:embedding medium and allow it to stand for at least 1 hour at room temperature. Remove the mixture, replace it with 100% embedding medium and leave for 6-12 hours at room temperature. Embedding Transfer each sample to a dry capsule or mould and fill the mould with embedding medium. Cure the medium in an oven at 60°C for 12 hours or until it is hard. Block hardness increases with curing time and temperature. If a curing oven is plugged into a lamp lighter/timer, this may be set to run at night and blocks will always receive the same curing time if placed into the oven during day hours and if they are removed any time on the following day. Araldite is very viscous and infiltrates poorly, but it stains and sections better than Spurr's resin. Blocks can be trimmed and sectioned after the blocks return to room temperature. References Finck, H. (1960), J. Biophys. Biochem. Cytol. 7, 27-30 Luft, J.H.(1961), J. Biophys. Biochem. Cytol. 9, 409-414 Glauert, A.(1991), Microscopy and Analysis, September; 15-20
Aurion Immuno Gold Reagents
*Published in AURION IMMUNOGOLD REAGENTS EDITION III https://aurion.nl/sharing-our-knowledge/ What kind of particle size should I use? Always use the smallest particle size to fit your application. Conjugates based on smaller particles are more efficient than larger particle based conjugates. If visualisation is difficult with smaller particles these can be enlarged with silver enhancement. Very sensitive specimens for SEM observation are best served with a larger particle size conjugate. This prevents ultra structural enhancement reagents. Is it true that gold conjugates are more background prone than other conjugates? No! This fairy tale comes from the fact that gold conjugates are based on particles and that visualisation is also based on separate particles. Contrary to enzyme and fluorescent markers, gold conjugates are more like a digital system, either they are there and then you will see them, or they are not present. Enzyme and fluorescent markers are sooner to be considered as “analogue” markers, their visibility in detection increases with their local concentration or with the time the enzyme marker can produce a visible reaction product. An unbiased look at controls in fluorescence shows always a low level of light that is inherent to the presence of double bonds in biological compounds and on top of this comes the fluorescence from the labeled antibodies. Likewise will an unbiased look at control specimens incubated only with alkaline phosphatase or peroxidase labeled antibodies usually show a faint overall staining of the specimen. Such faint levels are easily accepted or even filtered out. You cannot do this with gold conjugates since they are based on particles. Should I use a secondary gold conjugate or Protein A (or G)? That depends on what your goal is. Using secondary conjugates results in a higher labelling density. Therefore it is often said that secondary conjugates are more sensitive than Protein A conjugates. This is partly true. Protein A (or G) recognises only one site on a primary antibody molecule. Binding will occur only when this site is available and not obscured by its environment. Secondary conjugates recognise more sites on primaries and therefore the chance that a primary antibody will be detected is greater. Essentially this is the increase in sensitivity. If all primary antibodies would be available to the sane extent for binding to either Protein A or a secondary antibody conjugate, then the use of the latter would only result in more particles. This helps in localising antigens at low magnifications, in other words this is an increase in detectability. Is it possible to do pre-embedding labelling of intracellular antigens? Yes. Single cells are most suited. Plant material with a thick impenetrable wall is not. The Ultra-Small gold conjugates are the conjugates of choice. In many cases a permeabilization step with NaBH4 suffices to open up the specimens and allow penetration of reagents. Low concentrations of mild detergents like saponin help. One thing should be emphasised: reaction times have to be prolonged since full penetration of the reagents to the internal antigens has to be achieved. To remove unreacted reagents after incubation wash procedures have to be adapted likewise! The Aurion Newsletter #5 deals with this topic. How can I verify that my conjugates are still active? There is a simple procedure to check this. It is described in great detail in Aurion’s Newsletter #4. In short: you need a nitro-cellulose strip, apply dots from a dilution series of your primary antibody and incubate the strip with the gold reagent. The dots will stain red with the larger conjugates. When testing an Ultra-Small conjugate silver enhancement has to be applied for visualisation. How can I verify that the silver enhancement reagents are still fine? Again, there is a simple procedure to check this. It is described in great detail in our Newsletter #4. In short: you need a nitro-cellulose strip, apply dots from a dilution series of your gold conjugate and incubate the strip with the silver enhancement reagents. The dots should become brown-black. During this period of time the mix of reagents should remain glass clear without any visible presence of silver caused by auto nucleation. The activity of the Silver Enhancement reagent SE-EM for Electron microscopy can be tested by adding 10μl of the enhancement mix. The solution s should turn yellow in 30-45 minutes. Is it advisable to use outdated conjugates? As long as their reactivity is OK and there are not too many clusters formed this is no problem. Gold conjugates are very stable. There may be some release of protein from the particle surface with time, but generally this does not result in reduced reactivity. The reactivity of the conjugate is easily checked with a dot-spot test as described in Newsletter #4. Cluster formation may increase with time, depending on the type of conjugated protein and the particle size. The larger the particles the more clusters. These can be removed by centrifugation of the diluted conjugate before use. We have collected answers to frequently asked questions from immunogold users. They are listed below. If your question is not listed on this page or you have further queries, please contact us directly by phone or e-mail. We remain at your disposal. Is it possible to double label using two antibodies from the same animal source? Yes, there are ways to do this. One is by using Protein G or Protein A conjugates with different particle sizes. The procedure would be: first incubate with primary antibody l, detect this with Protein A (or G) with the smaller particle size. Then incorporate an incubation with excess free Protein A or G (50-100μg/ml). This will block practically all binding sites for Protein A or G. Next, incubate for the second antigen with primary antibody ll and detect this with the larger sized Protein A or G gold conjugate. A second possibility is to use one-step incubations with a mix of primary antibodies, each labeled directly with a different gold particle size. We offer a custom labeling service. Details can be found in the section on custom labelling. What kind of grids should I use for silver enhancement? Nickel is the material of choice. Gold grids are out of the question as they will be neatly enhanced as well. The same with copper. Nickel grids are preferred to copper ones for immuno incubations anyway, since nickel is more inert and less poisonous to immuno or enzyme reactions. Nickel grids can be annoying because of their magnetic properties. This is easily overcome by using either non-magnetic tweezers or by using a flattened loop to transfer grids from droplet to droplet during immuno incubations. We do sell a perfect-loop (EMS70944) for this application. What about silver enhancement and OsO4? OSO4 fixation can be used before incubation, after incubation or after silver enhancement. Because of its destructive effect on antigens OSO4 fixation is not often used when immuno incubations are intended. However, in general silver enhancing immuno incubated OSO4 fixed specimens causes no difficulties. An Osmium fixation step can be introduced after incubation to improve contrast in specimens. As stated before, applying silver enhancement generally causes no difficulties. On occasion in the past, using OSO4 fixation after silver enhancement used to lead to the removal of part of the deposited silver. However, with the introduction of SE-EMplus this is no longer the case, as the resulting enhanced particles are no longer sensitive to oxidation. I get no positive results, now what? When your incubated specimens look as clean as your controls, either (one or more of) the reagents are inactive, or the antigens are destroyed, masked or absent. The cause is easily found by performing tests working backwards through the incubation protocol using dot-spot tests as described in Newsletter #4. First test the activity of the silver enhancement reagents (if they were used at all) on the gold conjugate that was used. If silver enhancement is fine, the next step is to test the gold conjugate on the primary antibody used and so on. If it proves that the problem is not in the reagents, you will have to look into antigen preservation. Is a different fixation due? Or a different embedding medium? Using light microscopical evaluation of the results such questions are answered without tedious EM experimental work. I am having background problems. Is this due to the gold conjugate? When specimens are blocked correctly and the right composition and condition of incubation buffer is used, background levels should not be interfering with specific signals. Some background will always exist: to some extent all compounds have a certain affinity for other compounds and depending on availability and concentration an interaction may occur. There is no absolute black and white in this respect. When you leave out the primary antibody incubation and only use the gold step and your background has become much reduced, then your primary antibody causes background. Remedy: purify the primary antibody by either affinity chromatography (in case of an antiserum) and/or by cross-adsorption. If you have unacceptable levels of background without using a primary incubation, then the specimen has a tendency to bind to gold conjugates. Background may have many causes which are centered around three different types of interactions: Residual fixative activity, which is eliminated by using a NaBH4 or Glycine block step prior to the protein block step. Stickiness to hydrophobic areas (embedding medium, lipid rich specimen compounds). This is reduced by using an adequate protein block step involving a partly hydrophobic protein like BSA or Casein. Charge-based interactions causing negatively charged reagents such as antibodies and gold conjugates to adhere to oppositely charged areas in the specimen (notorious are the histone proteins, some collagen types and poly-L-lysine that is sometimes used to make sections stick to surfaces). This type of interaction can only be overcome by adding an excess of negatively charged indifferent molecules to the incubation media. We have developed a chemically modified BSA especially for this purpose. Newsletter #1 gives indepth information regarding it. Immunogold Silver Staining of E-cadherin on a paraffin section of human skin. Courtesy of R. Moella, Dept. of Exp. Path., EUR, The Netherlands. Mouse monoclonal anti E-cadherin GAM lgG UltraSmall Aurion R-Gent SE-LM Are there any fora which I can address with questions regarding labelling or microscopy? There are a few newsgroups which may be of interest: bionet.cellbiol, bionet.cellbiol.cytonet, bionet.molbio.methdsreagnts and sci.bio.immunocytochem. There is a microscopy listserver to which you can subscribe and which offers a platform to ask questions regarding light and electron microscopy in all its facets. You can subscribe by sending an e-mail message to ListServer@MSA.Microscopy.Com. The message only has to contain the words "subscribe microscopy". How can I do a controlled silver enhancement with pre-embedding? With pre-embedding there are 2 possibilities: either the enhancement is done before embedding or on the sections after embedding. We prefer to do the enhancement on sections (on nickel grids) since this gives more control over the degree of enhancement. Using longer enhancement times allows to observe larger (even ultra thin) sections in the light microscope. This facilitates searching for the area in the specimens where a reaction has occurred and allows easy targeting and trimming down to the area of interest for EM sectioning. Shorter enhancement is then used on sections for EM. Using enhancement before embedding has the disadvantage that once enhancement proves to be too long (resulting in too large particles) this can not be reversed. In which case should I use a Single Fab or F(ab’)2 conjugate instead of the complete immunoglobulin conjugate? The size of a conjugate is co-responsible for its efficiency. The overall size is determined by the particle size and by the size of the proteins adsorbed onto the particle surface. That is why we introduced UltraSmall particles in the first place. Whenever a specimen is relatively dense or intensely cross-linked immuno reagents will be more hindered in their action. If you are already using an Ultra-Small conjugate further improvement may result from using a single Fab or F(ab’)2 fragment of the specific secondary antibody instead of the intact lg-molecule. When should I use normal serum in the incubations? It is a good idea to use normal serum as an additive to the blocking and incubation buffer when using secondary antibody conjugates. The normal serum should be the same species as the secondary antibody conjugate. Its action is similar to the action of BSA. Please be careful when using normal sera to suppress background with Protein A or Protein G conjugates. These conjugates detect several lg-types from different species which, when used as normal serum additive, would lead to an impressive amount of gold particles all over the specimen. We offer several Blocking Solutions tailored for specific secondary antibody or protein A/G incubations. What about sensitivity, signal-to-noise ratios and detectability? Sensitivity can be considered at different levels in the total preparation and incubations. Ideally during preparation one would like to preserve all antigens present. In many cases this is not possible. But at least a representative fraction should be preserved and be available for immuno labeling. It all depends on the preparation procedure (fixation, embedding, temperature, etc.), which leaves you with a specimen or section with a given number of available and recognisable antigens. The ensuing detection protocol has 100% sensitivity if all the remaining antigens are detected, i.e. are represented by at least one gold particle or marker molecule. Again, due to masking and steric hindrance by the specimen composition this will only in exceptional cases by fully attained. The immuno labeling sensitivity thus expresses the degree to which available antigens can be detected by the employed combination of primary antibody and secondary conjugate. The quality of the primary antibody is the next important item. Theoretically the Kd-value of an antibody/antigen reaction is a measure for the dilution at which the incubations should be performed and for the stability of the ensuing bond. Sensitivity will go up with more concentrated antibody solutions up to a maximum level. However, when the primary antibody shows cross-reactivity there is not necessarily an improved signal-to-noise ratio. The reliability of the detection by the primary antibody improves in such cases with higher dilutions, probably leading to a smaller amount of antigens detected, but to an improved signal-to-noise ratio. Thus, sensitivity at the level of the primary antibody has to be balanced against the signal-to-noise ratio. The last step is the quality of the secondary reagent. In fact you will be looking at a number of gold particles which represents a number of secondary antibodies which have detected a number of primary antibodies. For the interaction between the secondary reagent and the primary antibody the same rules apply as indicated for the antigen/primary antibody reaction. Detectability reflects the degrees to which the final result of all the reactions involved can actually be seen. This is depending on the right match between particle size and magnification. Ultra-Small particle-based conjugates for instances are among the most efficient detection systems, but you will only detect them after silver enhancement (in most applications). What is epi-polarisation microscopy? Epi-polarisation is a technique used for the very sensitive light microscopical observation of metal particles. Where bright field microscopy depends on contrast levels in discriminating signals, epipolarisation works differently: provided particles are large enough individual particles will be observed. So in fact you are evaluating your labeling results on the same basis as with an electron microscope by looking at individual particles. This makes this technique so valuable as it builds a bridge between the light level and the electron microscopical observation. What you need to do is this: a high-quality light microscope equipped with an epi-illumination source, preferably a high pressure Hg-lamp (although a halogen source may also do). Many laboratories have an epi-fluorescence microscope at their disposition with a 40X (or higher) oil objective. Such microscope equipment forms the correct basis. You only need to implement an epi-polarisation filter (the so-called epi-block or IGSS filter) in the filter housing. The epi-block contains two polarisers, differing 90 degrees in orientation with respect to each other. How does it work (in short)? High intensity light passes the first polariz\ser in the epi-block and becomes polarised. The polarised incident light passes the objective lens and interacts with the specimen. The biological material hardly gives any reflection, and the reflected light is unmodified. The metal particles mirror the polarised light, thereby randomising the polarisation angle. Reflected light passes up through the objective lens. On its way to the eye pieces or the photo camera the light passes the second polariser in the epi-block. While doing so, light with the original polarisation angle (the way it was polarised in the first place before ever hitting the specimen) is extinguished, whereas light that has become randomly polarised (and which comes from the silver metal particles) passes the epi-block. As a result you will see individual bright stars (the gold/silver particles) against a dark background. Epi-polarisation observation can be combined in real time with bright field imaging, providing for a very sensitive detection of even extremely low amounts of antigen while still having the advantage of full morphological details in the specimen. My specimens for pre-embedding have a lot more antigens than a corresponding ultrathin section. Should I use more concentrated reagents? The increased amount of antigens should be balanced by a larger amount of reagent volume at an appropriate dilution (the same as used on sections with low amounts of antigen), and not by more concentrated reagents. The reason is that with increased concentration more cross-reactions may occur and signal-to-noise ratios will decrease. Incubating specimens for pre-embedding in larger volume quantities is best performed on a rocking table for a prolonged time to warrant penetration to antigenic sites in the specimen. What is the advantage of gold nanoparticle conjugation via adsorption? This is an easy and straight forward method for the conjugation of high molecular weight macromolecules. The flocculation test that tests the effect of a high salt concentration on the stability of the complex makes it easy to check if adsorption to the gold nanoparticle surface is successful or not. Can you give me more information on gold nanoparticle conjugation strategy? First check if conjugation via adsorption is feasible. If yes, check bio activity e.g., via a dot spot test. Biomolecules that are too small to be conjugated via the classic adsorption method can be covalently linked to functionalised gold nanoparticles as an alternative. A prerequisite for successful conjugation to Carboxyl-functionalised gold nanoparticles is the presence of primary amine(s) in the target molecule. As a rule of thumb conjugation via classic adsorption to conventional gold nanoparticles is possible with macromolecules having a MW > 40 kDa. What is the conjugation principle behind covalent conjugation using carboxyl-functionalised gold nanoparticles? The conjugation relies on the well known and proven EDC/sulpho-NHS chemistry. EDC/sulfo-NHS activation at pH 5 results in an amine reactive ester, immediately followed by binding to free amine on the target molecule. For more information and FAQ, please visit https://aurion.nl/sharing-our-knowledge/
Cargille Refractive Index Liquids
Cargille Refractive Index Liquids have become standard items in many laboratories as their applications have expanded from routine mineralogical identifications and quality control. New and broader uses in many more fields such as chemicals, engineering, medical, forensic, optics, and instrumentation are continuously discovered. Many special requirements for specific applications have created a need for more technical data, new formulations, extended ranges, smaller increments and higher degrees of precision. Since 1942, Cargille Laboratories, Inc. has been filling this growing need. And today, the largestand most comprehensive assemblage of refractive index liquids - over 250 stocked items - are available for science, industry, medicine, and education. For specialised and unusual applications, many more can be formulated and others are routinely researched in anticipation of new needs. Chlorofluorocarbon (CFC) components used in some Cargille optical liquids have physical properties that are harmless to the ozone layer, unlike those found in refrigerant gases, propellants, and solvents, which are destructive. The components used by Cargille Laboratories have vapour densities ten times that of air, are relatively nonvolatile, and have boiling points at least 100°C higher than the highest-boiling CFC of those listed for removal from commercial use by the Montreal Protocol. Applications Cargille Refractive Index Liquids have a broad range of applications in diversified fields. The following table briefly illustrates some of the typical areas of use. AREA OF INTEREST TYPICAL APPLICATIONS IDENTIFICATION Identify minerals, ores, chemicals, specimen fragments, plastics, gems, translucent or transparent solids by microscopic immersion techniques. MOUNTING MEDIA Temporarily mount specimens in various index media for matching or contrasting index combinations. Mount specimens and thin sections in a stable, viscous, nondrying index of refraction liquid to permit sample rotation by shifting cover glass for more comprehensive examination. (Cargille MeltMount) OPTICAL ANALYSIS Microscopically study solids by dispersion staining, focal masking, and double variation refractometry techniques with high dispersion liquids. REFRACTOMETRY Calibrate refractometers and other optical instruments. EDUCATION Instruct and exhibit principles and applications of refraction. SPECTROPHOTOMETRY Determine refractive index of specimens by referral to calibration curves relating wavelength to index of refraction. STRAIN ANALYSIS Examine stress and strain effects of transparent or translucent items; moulded, formed, curved or intrinsically shaped parts by polariscopic immersion technique. (Cargille Immersion Liquids) TOXEMIA Identify salts precipitated from body fluids and measure relative amounts of sodium and potassium chloride for indicating toxemia. OPTICAL COUPLING Couple optical elements with liquids formulated for refractive index and viscosity. OPTICAL LENSES Obtain unique optical properties of hollow lenses by filling with index of refraction liquids. ELECTRO-OPTICS Examine and preserve cathode coating without stripping or re-immersion by utilising oils matching index of crystals. POLLUTION Identify particles and particulates from air or water. FLUID FLOW Photograph flow patterns by filling test system with refractive index liquids having suspended "beads". Refractive Index Liquids: Series A nD 1.460-1.640 ±0.0002 (589.3nm, 25.0°C) Series AA nD 1.400-1.458 ±0.0002 (589.3nm, 25.0°C) Series AAA nD 1.300-1.395 ±0.0002 (589.3nm, 25.0°C) Series B nD 1.642-1.700 ±0.0002 (589.3nm, 25.0°C) Series E nD 1.500-1.640 ±0.0005 (589.3nm, 25.0°C) Series M nD 1.705-1.800 ±0.0005 (589.3nm, 25.0°C)
CCC Carbon adhesive (DG)
CCC is an electrically conductive cement for specimen mounting in SEM work, featuring good adhesive quality. Due to its electrical conductivity, all conductive specimens are ready for investigating, immediately after drying of the cement. Nonconductive specimens must be coated with carbon or metal, but no additional treatment is necessary such as painting conductive bridges, which may contaminate the specimen by spreading of silverdag or solvent. CCC contains only hydrogen, carbon and oxygen with no other residue after drying. For that reason, no characteristic spectral peaks are found in CCC. When analysed with an energy dispersive spectrometer, there is only the continuum to be found. If the entire specimen holder is painted with CCC, the cement will mask the holder and there are no X-rays produced in the holder, which could interfere with the analysis of the specimen itself. Application: For powders or very light specimens, a thin layer or for bulk specimens, a heavier layer is painted on the specimen stub. If powders are to be mounted, allow a short drying period (of several minutes), so that the powder does not sink into the wet glue. Trial and error will show the correct predrying. Heavier specimens can be placed directly into the wet paint. After drying, only carbon (with acrylic binding agents) is left. Drying is simply the physical process of evaporation of the organic solvents. Depending upon the thickness of the layer, the drying time varies from a few seconds (very thin film) up to 30 minutes (for a heavy coating). The rate of evaporation is set in such way, that the entire bottle can be used up without thickening of the contents - assumed closing the bottle after each use. If however the viscosity does get too high, CCC Thinner may be added. Xylene or toluene easily cleans tools or instruments which accidentally were contaminated with CCC.
Citifluor™ Antifadent Solutions
Mountant Media Containing Antifadents – The Antidote For Photobleaching What is an Antifadent Solution? Antifadent Solutions reduce the photobleaching or fading of the fluorescence of dyes used for labelling biological species. The fading of fluorochrome dyes is a particular problem in fluorescence microscopy, such as in immunofluorescence studies. Fluorescent dyes are also used as cell markers, for following the uptake and release of calcium from cells for example, and for characterising cell surfaces. Non-fluorescent immersion oil is available for work at high magnifications. In fluorescence microscopy, the fluorescence is stimulated by high intensity UV or visible light. Absorption of the light populates an excited state of the dye (usually a singlet state) and this leads to fluorescence. However, the excited state of the dye may undergo chemical reactions which leads to its destruction as evidenced by the fading or bleaching of the fluorescence and consequent loss of the image or in the case of assays, a change in signal intensity during measurement. Using antifadent (anti-bleaching) solutions as mounting media are ideal to overcome the problems of examining samples by fluorescent microscopy or other detection systems and as an additives for assays. The most frequently used solution is AF1 which is an antifadent (anti-bleaching agent) contained in a glycerol PBS (phosphate-buffered saline) solution and is particularly useful for examining tissue sections. AF2 solution contains the antifadent in glycerol which enables users to choose their own buffer, and AF3 is the antifadent in PBS solution. AF3 is particularly useful for examining live cells. AF87 is a non-fluorescent immersion oil containing antifadent. CFPVOH is an aqueous solution of poly (vinyl alcohol) for use as a solid mountant. AF100 is a solution of antifadents (anti bleaching agents) for use with CFPVOH when fading (bleaching) of specimens in a solid mountant is a problem. These antifadents are not based on p-phenylenediamine. All solutions may be stored at room temperature. Choosing the Best Antifadent These are the critical decision factors which will help you select the most appropriate mountant: Do you need a Hardening or a Non Hardening medium? If your system is Glycerol tolerant than choose a Glycerol Based product, if not then use a Glycerol Free solution These high refractive index mountant solutions are to be used where you wish to obviate the effects of spherical aberration (caused by a mismatch between the refractive index of the glass of the coverslip and the mountant medium) which leads to a loss of resolution of your images. Table 1. Types of antifadent available OVERVIEW OF APPLICATIONS Non-Hardening Mountants: Use of Non-Hardening Citifluor antifadents Glycerol Based AF1 A glycerol-PBS solution contains an amine antifadent AF1 plus DAPI contains both an amine antifadent and the DNA stain, DAPI AF2 A glycerol solution contains an amine antifadent that allows you to choose your own buffer solution AF4 A glycerol solution of n-propyl gallate High Refractive Index Glycerol Based Solutions These solutions have a refractive index that matches that of the glass of the coverslip thereby minimising the effects of spherical aberration. Such mountants are invaluable for imaging the internal structures of samples by techniques such as single multi-photon confocal niche since it acts as an efficient Clearing Solution (clearing occurs within a few minutes – Sean Speese, Oregon Health and State University, Personal Communication). The beneficial effects of these reagents can be seen in the two figures below where high resolution images have been obtained at a significant depth within the mountant solution. Additional features of these solutions includes their total water miscibility, they are non-odorous and do not quench the fluorescence of fluorochromes (unlike thiodiethanol, another proposed high refractive index mountant) and the solutions are stable for long periods (at least 1 year). CFM-1 is a glycerol-PBS buffered solution having a refractive index of ~1.52 which can be used for transmission microscopy as well as with epifluorescence microscopy CFM-1 plus AF is a glycerol-based solution having a refractive index of ~1.52 (at room temperature) that contains an amine antifadent CFM-2 is a glycerol-tris amine buffered solution having a refractive index of ~1.52 (at room temperature) and a pH of ~8.5 CFM-3 – This glycerol-based antifadent contains a phoenlic antifadent of neutral pH and a refractive index of ~1.52 which also acts as a Clearing Solution enabling visualization of fluorochromes deep within the sample CFMR2 –This product was designed for use with samples labeled with GFP. It contains a unique antifadent which does not de-oxygenate the solution which is an essential property if the GFP is not to bleach Glycerol Free AF3 is a PBS solution containing an amine-based antifadent CFPVOH is an aqueous solution of poly (vinyl alcohol). Provided care is taken not to allow the water to evaporate, it will remain fluid. Retardation of photobleaching is effected by the addition of the aqueous antifadent solution AF100 AF100 Is a PBS solution of an amine-based antifadent that if used as an additive to CFPVOH (1 part AF100 to 9 parts CFPVOH). Provided care is taken not to allow the water to evaporate, it will remain fluid. AFR3 is a PBS solution containing a NEW non-amine, non phenolic antifadent High Refractive Index Glycerol Free Solutions AF87 is an immersion oil having a refractive index of 1.52 and contains an antifadent. It may be used as an immersion oil and also as a mountant. Since AF87 is an oil that is immiscible with water, it is essential that specimens are dehydrated before application of the mountant. Hardening Mountants: Hardening mountants are aqueous solutions containing a polymer such a poly (vinyl alcohol) (PVOH). When a few microlitres of these solutions are pipetted onto a microscope slide and a coverslip applied, evaporation of water slowly takes place and the formation of a stable film results which immobilises the coverslip. A variety of hardening mountants is available which provide films possessing a range of hardness. They are based on water-soluble polymers such as poly(vinyl alcohol(PVOH) and poly (vinyl pyrrolidone)(PVP). Why are aqueous glycerol solutions of PVOH containing antifadents inherently unstable? PVOH is produced by hydrolysis of poly(vinyl acetate) and most commercial samples of PVOH contain residual (unhydrolysed) acetate groups. These groups undergo hydrolysis, often accelerated by the added antifadents, during storage and this can cause a change in pH and more usually gellation. This latter process leads to an unpredictable shelf-life. By making up small volumes of PVOH solution containing the required amount of antifadent solution, you have materials of consistent composition and performance as well as making better use of your purchased materials. Glycerol Based AF200 is a glycerol solution containing an amine-based antifadent AF300 is a glycerol solution containing a phenolic type of antifadent Film Forming Polymer Solutions Tris-MWL 4-88 is a classical, popular mountant solution based on Mowiol® 4-88, glycerol, water and tris-amine buffer. Following evaporation of the water a film of weak to medium strength is formed. To have effective reduction in photobleaching, it should be used with either AF100, AF200 or AF300 CFPVOH is an aqueous solution of poly (vinyl alcohol) and is designed to be used with the glycerol-based antifadent solutions AF200 or AF300. Following evaporation of the water films of medium hardness are produced Glycerol Free PVP plus antifadent is an aqueous solution of poly (vinyl pyrrolidone) containing an amine-based antifadent. The solutions are stable over long periods e.g. in excess of five years CFPVOH plus antifadent is an aqueous solution of a carefully selected PVOH containing an amine-based antifadent and has a shelf life 6 months Film Forming Polymer Solutions CFPVOH is an aqueous solution of poly (vinyl alcohol). Retardation of photobleaching is effected by the addition of the aqueous antifadent solution AF100. (1 part AF100 to 9 parts CFPVOH)
Citifluor™ Non-Hardening Antifadent Solutions
EMS17970, EMS17971, EMS17973 Glycerol Based High Refractive Index Glycerol Free High Refractive Index AF1 CFM-1 CFPVOH + AF100 AF87 AF1 + DAPI CFM1 + AF AF3 AF2 CFM-2 AFR3 CFMR2 AF4 CFM-3 Glycerol Based Antifade Reagents Antifadent pH RI P/G* AF1 ~9 1.463 P AF2 NA 1.473 P AF4 NA 1.476 G * P = HDPE plastic bottle, G = Brown glass bottle AF1 Usage AF1 is a mountant solution composed of glycerol, phosphate buffered saline and an antifadent. It was specifically designed to stop the photobleaching of the fluorescein moiety of FITC labelled biological specimens. AF1 is useful for many other fluorochromes such as DAPI, rhodamines, Hoechst, Alexa and cyanine (Cy-3 and Cy-5) dyes, Texas Red, phycoerythrins and Green Fluorescent Protein (GFP). It is ideal for examining tissue sections and dead cells. In addition, it has been found useful for stabilising the AUTOFLUORESCENCE of species such as cyanobacteria. AF1 solutions have been employed with the following techniques; fluorescence in situ hybridisation FISH (including CARD-FISH) and confocal laser scanning microscopy (CLSM) are being used. Applications This well-established product, has an anti-bleaching (anti-fadent) contained in glycerol-PBS (phosphate-buffered saline) solution and was specifically designed to alleviate the PHOTOBLEACHING of antibodies labeled with FITC (1-4). AF1 Example A AF1 is useful for many other fluorochromes such as DAPI (1, 4, 5-10), rhodamines (11-17), Hoechst (11, 18-24), Alexa (9, 12, 25-29) and cyanine (Cy-3 and Cy-5) (30-37) dyes, Texas Red (4, 38-42)), phycoerythrins (43-48) and Green Fluorescent Protein (GFP) (7, 26, 39, 41, 49-54). It is ideal for examining tissue sections and dead cells. In addition, it has been found useful for stabilising the AUTOFLUORESCENCE (46, 55-57) of species such as cyanobacteria. AF1 Example B Storage and Shelf-Life AF1 solutions may be stored at room temperature and ideally between 5° and 15°C and out of strong sunlight. The cap of the bottle or if using the pipette supplied with the material, the cap which covers the pipette delivery point, should always be replaced after use as a matter of good practice. Samples stored under these conditions for 6 months have shown no apparent deterioration. If the AF1 solution is being used in an assay, a control experiment should always be carried out. Useful tips In some cases, the reduction in the rate of PHOTOBLEACHING may be accompanied by a reduction in the initial intensity of the fluorescence signal. By diluting the AF1 solution with glycerol the reduction in the intensity of the fluorescence signal can oftentimes be mitigated. Dilution of AF1 with glycerol will increase its refractive index whereas dilution with water will reduce its refractive index. AF1 solutions have been employed where the techniques of fluorescence hybridization (FISH) (5-7, 9, 11, 17, 23, 28, 34-37, 58-60, 62-66) including CARD-FISH (63-65) and confocal laser scanning microscopy (CLSM) (1, 2, 29, 30, 36, 39, 50, 52, 54, 67, 68, 69) are being used. AF1 Example C The solution has a pH of ~10, a refractive index of 1.463 at 20°C and is optically transparent from 300nm into the 750nm. The solution should be pipetted (15 to 25µl) onto the specimen and then a cover slip applied. Specimens mounted in this way may be kept in a refrigerator without having to seal the edges of the coverslip with a material such as nail varnish, and usually they retain their fluorescence for many months. Obtaining the correct viscosity for your application The solution is of medium viscosity and has a water-white in appearance. It may be stored at room temperature and ideally between 5° and 15° and out of strong sunlight. The cap of the bottle or if using the pipette supplied with the material, the cap which covers the pipette delivery point, should always be replaced after use since the solution is hygroscopic. Samples stored under these conditions for 6 months have shown no apparent deterioration. If the AF1 solution is being used in an assay, a control experiment should always be carried out. If the viscosity of the AF1 solution is too high for your purposes, it may be admixed with AF3 mountant (a PBS solution of the same antifadent as is present in AF1) solution. As the amount of AF3 solution is increased so the viscosity decreases. Conversely, if you wish to have a higher viscosity add AF2 mountant solution to the AF1 solution. Graph 1. Viscosity of AF1 solution when AF3 solution is added AF1 can be used to make up a hardening mountant. To a poly (vinyl alcohol), e.g. Airvol 203 (Air products) or Mowiol® 4-88 (Calbiochem) solution (20% in water), add ~20% by volume of AF1. This solution is best used soon after it is prepared as it doesn't have a long shelf-life AF1 plus DAPI Mountant Solution This is a new addition to the range of Citifluor products and is composed of an AF1 mountant solution to which DAPI has been added at a concentration of 2µg/ml. If a lower concentration of DAPI is required, dilution should be carried out by adding a further quantity of AF1 solution. The solution will be of value to those using the FISH technique. It is recommended that the solution be kept in a tightly sealed brown glass bottle and stored in a refrigerator at a temperature between 0° and 5°C. AF2 Properties and method of use AF2 Example A AF2 is a mountant solution composed of glycerol and an antifadent and is optically transparent from 300nm into the 750nm. It was specifically designed to stop the photobleaching of the fluorescein moiety of FITC (70-73) labeled biological specimens. AF2 Example B Its application is not however limited to FITC labeled materials and has been used with advantage with many other fluorochromes including rhodamines (71), DAPI (71, 73, 74-78) and GFP (74, 80-82). It is ideal for examining tissue sections and dead cells. An aqueous solution (75% AF2 to 25% water v/v) has a pH of ~10. AF2 solutions have been employed where the techniques of fluorescence in situ hybridization (FISH)(73, 76, 77, 79, 83), and confocal laser scanning microscopy (CLSM) (70, 71, 73, 75, 79) are being used. Storage and shelf-life The solutions are of medium viscosity and are water-white in appearance. They may be stored at room temperature and ideally between 5° and 15°C and out of strong sunlight. The cap of the bottle or if using the pipette supplied with the material, the cap which covers the pipette delivery point, should always be replaced after use as a matter of good practice and also to prevent the ingress of water (due to the glycerol being hygroscopic). Samples stored under these conditions for 6 months have shown no apparent deterioration. If the AF2 solution is being used in an assay, a control experiment should always be carried out. Useful tips The viscosity of AF2 solution is influenced by adding AF1 solution. Graph 2. Viscosity of AF2 solution when AF1 solution is added If the viscosity of the AF2 solution is too high for your purpose, it may be reduced by the addition of either AF1 or AF3 solutions. The AF2 mountant solution should be pipetted (15 to 25µl) onto the specimen and then a cover slip applied. If the slides are stored in a refrigerator, the viscosity of the mountant solution increases thereby helping too keep the cover slip in place. There is no need to seal the cover slip with nail varnish. Specimens mounted in AF2 solutions have been kept in this way for many months without loss of fluorescence intensity. F2 has also been used to create a hardening formulation (84) by adding it to an aqueous solution of poly (vinyl alcohol), e.g. Airvol 203 or Mowiol® 4-88. AF4 Properties and method of use AF4 is a solution composed of the antifade reagent n-propyl gallate (85) dissolved in glycerol and is optically transparent between 400 nm and 750nm. If the solution is to be used as a mountant solution as supplied, wash the specimen with a buffer of appropriate pH for the fluorochrome label (so as to maximise the fluorescence intensity) and then apply a few drops of AF4. Alternatively, buffer solutions may be added to AF4 to create your own mountant solution but it is inadvisable to make solutions containing buffer in excess of 30% by volume since this leads to a dramatic reduction in shelf-life. N-Propyl gallate is recommended in particular when DAPI is being as the fluorochrome. Storage and shelf-life The mountant solution should be stored in brown glass bottles at temperatures between 5° and 15°C and shielded from strong sunlight. The cap of the bottle should always be replaced after use as a matter of good practice and to prevent the absorption of moisture from the atmosphere. Samples stored under these conditions for 12 months, have shown no apparent deterioration and in particular no discolouration. If the AF4 solution is being used in an assay, a control experiment should always be carried out. Its application is not however limited to DAPI labelled materials and has been used with advantage with many other fluorochromes including fluorosceins, Alexa and Hoechst dyes and rhodamines. It is ideal for examining tissue sections and dead cells. The solution should be pipetted (15 to 25µl) onto the specimen and then a cover slip applied. If the slides are stored in a refrigerator, the viscosity of the mountant solution increases thereby helping too keep the cover slip in place. There is no need to seal the cover slip with nail varnish. Glycerol Based High Refractive Index Solutions Reagents for Transmission Microscopy as well as Fluorescence Microscopy. All the following solutions have a refractive index of ~1.52 at room temperature (20°C) and are freely miscible with water. They may be used as an immersion oil in the standard way as well as a mountant solution. The refractive index of the materials makes them ideal for use when specimens are being examined by confocal laser scanning microscopy (CLSM)). They show very little absorption above 400nm. The CFM range of mountant solutions should only be used with samples that have been fixed (4% paraformaldehyde in phosphate buffered saline for 30 minutes). The crosslinking (fixing) of the sample prevents non-covalently bonded fluorochromes, e.g. DAPI, Hoechst dyes, from becoming detached from the specimen etc. If some bleeding of the dye(s) is still observed a further fixation with paraformaldehyde should be carried out. For sealing the coverslips, when CFM mountant solutions are used, the product CoverGrip® marketed by Bioutium has been recommended. (We are grateful to Dr. Sean Speese, Jungers Center for Neuroscience, Oregon, U.S.A.for these many helpful suggestions). The two images below are examples of Citifluor product applications, kindly provided by Dr. Sean Speese: Citifluor product application examples Above: 63x1.4 NA PlanApo / 1 Airy / Nyquist sampling xyz / Zeiss LSM 710 Genetically expressed GFP-tagged nuclear protein in brain slice. Right: 3D rendering of nuclei stained for LamDm0 in Drosophila tissue, mounted in CFM3 + antifade, imaged via confocal microscopy. Notice the staining intensity is consistent throughout the entire 55µm stack with no increase in spherical aberration. CFM-1 This glycerol-phosphate buffered saline based solution has been specially formulated so as to have a refractive index of ~1.52 (at room temperature). The solutions are of medium viscosity, are water-white in appearance and have a pH of ~7.5 and may be used for conventional microscopy (transmission microscopy) as well fluorescence microscopy. Storage and shelf-life The solution may be stored at room temperature. The cap of the bottle should always be replaced after use. Samples stored under these conditions for 6 months have been found to exhibit little apparent deterioration. A few drops of the solution should be applied to the specimen, followed by a cover slip. An immersion oil (which may be a CFM solution if its viscosity is appropriate) is applied on top of the coverslip in the usual way. CFM-1 Plus Antifadent This glycerol-phosphate buffered saline based mountant solution has been specially formulated so as to have a refractive index of ~1.52 (at room temperature) and contains an antifadent to retard the bleaching of fluorochromes. The solution is of medium viscosity, are water-white in appearance and has a pH of ~9. Storage and shelf-life The solution may be stored at room temperature. The cap of the bottle should always be replaced after use. Samples stored under these conditions for 6 months have been found to exhibit little apparent deterioration. A few drops of the solution should be applied to the specimen followed by a cover slip. An immersion oil (which may be a CFM solution if its viscosity is appropriate) is applied on top of the coverslip in the usual way. CFM-2 A few drops of the solution should be applied to the specimen followed by a cover slip. An immersion oil (which may be a CFM solution if its viscosity is appropriate) is applied on top of the coverslip in the usual way. Storage and shelf-life This glycerol-tris-amine buffered mountant solution, has been specially formulated so as to have a refractive index of ~1.52 (at room temperature). It has a pH of ~8.5 which is appropriate for fluorescein conjugates. The solution may be stored at room temperature. The cap of the bottle should always be replaced after use. Samples stored under these conditions for 6 months have been found to exhibit little apparent deterioration. CFMR2 – A high refractive index mountant especially designed for samples labelled with GFP The Citifluor CFM series have proved very useful for imaging materials using CLSM. Samples labelled with GFP pose a special problem since the retention of a good image during imaging and upon storage is dependent upon the mountant solution containing oxygen. The classical antifadents consume oxygen when the fluorochromes are irradiated and in the case of GFP this accelerates the loss of the image. CFMR2 contains a unique antifadent that does not de-oxygenate solution but nevertheless, affords protection to the GFP. As with the other Citifluor CFM solutions the mountant has a similar refractive index to the biological tissue and hence enables a good depth of viewing within the sample. CFM-3 This antifadent is useful for retarding the photobleaching of DAPI, Alexa, Hoechst and cyanine dyes. CFM-3 Example A few drops of the solution should be applied to the specimen, followed by a cover slip. An immersion oil (which may be a CFM solution if its viscosity is appropriate) is applied on top of the coverslip in the usual way. Storage and shelf-life This glycerol-phosphate buffered saline based mountant solution has been specially formulated so as to have a refractive index of ~1.52 (at room temperature) and contains a phenolic type antifadent to retard the bleaching of fluorochromes. The solution is of medium viscosity, is water-white in appearance and has a pH of ~6.5. The solution may be stored at room temperature. The cap of the bottle should always be replaced after use. Samples stored under these conditions for 6 months have been found to exhibit little apparent deterioration. AF3 This well-established product was specifically designed to alleviate the PHOTOBLEACHING of specimens labeled with FITC (86,87). AF3 is useful for reducing the photobleaching of many other fluorochromes such as DAPI (88-93), rhodamines (94) and Alexa dyes (88, 91,95). It has been found useful for examining live cells although it should be noted that the antifadent may lead to cell lysis. Storage and shelf-life AF3 has an anti-bleaching (antifadent) contained in PBS (phosphate-buffered saline) solution. The solution has a pH of ~10, a refractive index of 1.338 at 20°C and is optically transparent from 300 nm into the 750nm. AF3 solutions have been employed where the techniques of fluorescence in situ hybridization (FISH) (86, 88, 89, 90, 97) and confocal laser scanning microscopy (CLSM) (88, 89, 90, 91, 93) are being used. The low refractive index of AF3 makes it ideal for use with the recently introduced technique of Total Internal Reflectance Spectroscopy (TIRF), where a large difference in refractive index between glass and the mounting medium is required (98, 99). The solution should be pipetted (15 to 25µl) onto the specimen and then a cover slip applied. AFR3 A new antifadent solution for use with live cells. The imaging of live cells in conjunction with an antifadent poses problems. The classical antifadents, amines and phenols can interact with the cell surface leading to lysis. The new solution contains an antifadent at a much lower concentration than that used is in the currently available solutions and is designed to show minimal phototoxicity. Unlike other antifadents it does not de-oxygenate the solution. The antifadent is supplied in a phosphate buffered saline solution. Texas Red, Alexa and Cyanine dyes have been stabilised with the new antifadent. This solution has a refractive index of 1.34 and hence is ideally suited for use with Total Internal Reflectance Microscopy (TIRFM). CFPVOH plus AF100 Although this combination of reagents is designed for use as a hardening mountant system (solid mountant system), it may be used in liquid form provided the water is not allowed to evaporate. A few drops of the solution should be applied to the specimen followed by a cover slip. Evaporation of the water under these circumstances takes a few hours. Storage and shelf-life The CFPVOH is an aqueous solution of poly (vinyl alcohol) and the AF100 is a phosphate-buffered saline solution of an antifadent. Solutions should be made up by mixing 1 part by volume of AF100 with 9 parts by volume of CFPVOH. These solutions should be used within 10 hours as the efficacy of the antifadent reduces with time. With this in mind, it is better to make up the mixture prior to use and not to rely on keeping solutions. AF87 This product is an immersion oil of low fluorescence that contains an antifadent. It is immiscible with water, is of medium viscosity and has a refractive index of ~1.52 at room temperature (20°C). This high refractive index makes it ideal for when specimens are being examined by confocal laser scanning microscopy (CLSM) (100). For use as a mountant, it is necessary to dry (dehydrate) the specimen before applying the AF87. Dehydration may be carried out by washing the specimen with ethanol, followed by drying in air (101, 102). AF87 has been found useful for examining specimens generated using the technique of FISH, labelled with the fluorochromes DAPI (103-105) and Cy dyes (101, 103). Bottles of AF87 should be stored at room temperature with temperatures of 50°C or lower being avoided since the low temperature may cause the development of a sediment in the bottle and a reduced antifadent performance.
Clear-Mount Aqueous Mounting Medium
IM032 - A direct substitute for Biomeda™ Crystal/Mount Description Clear-Mount medium is an aqueous mounting medium designed for permanent mounting of tissue sections and cell smears with peroxidase and alkaline phosphatase chromogens that can not be dehydrated with organic solvents. This mounting medium preserves Fast red, Aminoethylycarbazole (AEC), NBT/BCIP, INT/BCIP chromogens and is also compatible with counterstains like Haematoxylin and Nuclear fast red (NFR). It is also suitable for chromogens like DAB and DAB with nickel and cobalt. It is not compatible with H and E staining. Intended use Mounting of immunohisto slides. Reagent Ready to use mounting medium Storage 2 - 8°C, DO NOT FREEZE. Procedure Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 3-4 drops of mounting medium directly on top of the specimen and spread out evenly by tilting the slide back and forth or spread evenly with a 0.2 mL plastic pipette tip, making sure the tissue is not touched. Excess medium can be removed by touching the edges of slide against a paper towel. Let stand at room temperature for about 1 to 2 hours. Or the slides can be heated at 37-40°C for 40-60 minutes. The slides are ready for visualisation under a microscope. To visualize slide under microscope with 4X, 10X, 20X or 40X lens coverslip is NOT required For oil immersion lenses, a drop of oil can be placed directly on dried Clear-Mount medium for visualisation; the oil can be removed by wiping with Kimwipes. Handling and Storage This product is stored at room temp, however for long term storage 2-8°C is recommended. The pH of this product is more stable at 2-8°C. DO NOT FREEZE. Removal of coverslip If the coverslip has to be removed, soak the slide in warm (37°C) water for 5-10 minutes until the mounting medium is dissolved. Rinse slide with warm water to remove all mounting medium. The slide can be mounted again if required.
Coat-Quick 'G' Pen
To start, press the tip of the Coat-Quick "G" Pen straight down several times or until the fluid starts to soak the tip. Keep tightly capped and store cap side up. EASY TO APPLY: Place grids dull side up on #50 Whatman hardened filter paper and then just touch the pen tip to the dull side of the grid. Let sit for 1 to 2 minutes before using. Grids can be coated ahead of time and used later that day. Provides superior attachment of thin sections to the grid: When picking up sections from above, sections will adhere to the grid and not "pop" off when the grid is raised. This allows for reliable positioning of sections on the grid with minimal wrinkling or folding. During the staining process, the sections continue to adhere tightly to the grid, enabling one to thoroughly wash the grids without the sections lifting at the edges and folding over themselves. In the microscope, section crawl is minimised, since the sections are attached to the grid bars.
Colloidal Silver Liquid
EMS12630 EMS Silver Liquid is a unique air-drying silver used to convert a variety of surfaces, such as those electrically and thermally non-conductive to conductive for many application purposes. These silver liquids may be utilized during circuit repair processes or as RF shielding materials. During the application process, whether the user wishes to brush, spray, or dip the silver liquid, the material will harden immediately as the solvent begins to evaporate. We suggest that for optimal results across electrical and mechanical applications, the user cures the silver liquid at room temperature for approximately 16-20 hours or at 120-200°C for 30 minutes. The use of a heat gun is recommended to facilitate this curing process. Depending on the application of choice, one may consider applying additional layers of the silver liquid in order to reach an increased level of conductivity. Key benefits Item includes brush applicator Resistant to solvents Cures at room temperature Adheres to a variety of materials Electrically and thermally conductive Protocol Preparation of surfaces prior to application is optional, though not required Materials adhere to the following surfaces: boards that are polymer-based (phenolic), glasses, metals, ceramics, fiberglass, and many plastics Specifications Curing conditions Hardens instantaneously as the solvent evaporates % Solids 60% ± 1% Ag Sheet resistance 0.02-0.05 ohms/sq/mil (25µm) Thinner Catalog EMS12641 Silver Colloidal Extender Shelf life 6 months; for long-lasting results we highly recommend that the jar be rolled at 1-6 rev/hr. Note: Exposure to extreme temperatures is inadvisable in all circumstances. Storage Store the Liquid Silver in a dry location (we recommend between 5-30°C). Before use, make sure that the product is at room temperature. The product must be mixed completely to provide equal consistency.
Conductive Graphite Paint, SEM Adhesive 154 (DG)
Graphite Conductive Adhesive 154 is a graphite dry film lubricant that provides a clean, long-lasting lubrication without alteration of the dimensions of the component. This product requires simple surface preparation and can be applied to the majority of substrates with the use of spray, brush, or dip techniques. Not only is our product a significantly stable compound of processed micrographite and thermoplastic resin in isopropyl alcohol, but it also dries at room temperature. The Graphite Conductive Adhesive 154 requires minimal pre-treatment, is easy to apply, has a high lubricity, anti-seizes in nuclear applications, is applied as a concentrate, and has outstanding adhesion to most substrates. Instructions Surface Preparation Before coating substrates, we recommend that they be clean and dry. You may use a solvent wipe. For maximum adhesion, please note the following pretreatments: Mixing Our product is supplied in its concentrated form. Please note that it is thixotropic in nature and has the tendency to gel on standing. Preparation includes thorough agitation, then dilution with isopropanol to yield the required consistency for the application of choice. Application Diluted Graphite Conductive Adhesive 154 may be applied with the use of a spray, brush, dip, or roller. We do recommend using a spray, as it yields the most uniform coverage. Curing This product air dries within five minutes under normal temperature and humidity. Complete cure properties will develop in about two hours with consideration of film thickness and drying conditions. Technical Information Composition Properties Lubricant colloidal graphite Binder thermoplastic resin Fluid component isopropyl alcohols, esters, ketones Color black Solids content by weight 20% Consistency gel Density 0.9kg/L Flash point 10°C Tag closed cup Theoretical coverage 11.03m2/L @ 12.7µm VOC (dehydrates neat) 710g/L
Conductive Silver Liquid
Conductive silver liquid: Air drying Thinner: Methyl Ethyl Ketone or Toluene Application: Apply from nozzle, by brushing or dipping Shake well before use Keep tightly closed Store in cool, dry area. Refrigeration extends shelf-life This is a thermosetting material above 85°C. The solvent is "plumber's glue solvent" and available at hardware and plumbing supply places. WARNING: Flammable. Do not breathe fumes. Keep away from fire, heat. Re-suspending Silver Liquid The silver powder used is very small and affects the diffraction of visible light – so it no longer appears silver in colour. However, it is not nearly small enough, considering its high specific gravity, to remain suspended by Brownian motion. The silver powder settles fairly hard within a few days. Use a metal spatula to make it uniform. After that initial stirring, stir or simply invert every few days to maintain the suspension. Conductivity depends on the distance of silver particles from each other; at high concentrations the gap will be uniformly smaller. At some point with the gap too large, conductivity ceases. So stirring is important. Do not freeze, but refrigeration will extend the life of the binder.
Crystalbond™ and Wafer-Mount™
Crystalbond™ and Wafer-Mount™ wash away adhesives are ideal materials for temporarily mounting products that require dicing, polishing, and other machining processes. These adhesives exhibit high bond strength and adhere readily to metals, glass and ceramics. When processing is complete, Crystalbond™ and Wafer-Mount™ adhesives are easily removed by applying heat and cleaning with the appropriate solvent. Typical applications include: Matching advanced ceramics. Lapping and polishing optical components. Mounting cross-sections for electron microscopy. Dicing alumina and aluminium nitride ceramic substrates. Dicing germanium and silicon semiconductor wafers. Dicing ferrites and LCD glass. Dicing metallurgical specimens. Dicing metal and optical single crystals. Dicing piezoelectric transducers. Back-filling components for temporary mechanical support. Product Specifications: Crystalbond™ Product Crystalbond™ 509 Crystalbond™ 555 Crystalbond™ 590 Description Excellent adhesion to metals, glass, and ceramic. Transparent in thin cross sections. Minimises clogging of diamond tools. Low melting point adhesive. Soluble in hot water. Good for low shear processes. Transparent in thin cross sections. Excellent adhesion for cutting sub-miniature parts. Slightly flexible. Soluble in methanol or non hazardous 590-S stripper. Form Stick Stick Stick Size 7/8"x7" ½"x7"x1" 5/8"x1-¼"x7½" Weight 0.2 lbs/stick 0.15 lb/stick 0.5 lb/stick Flow Point 77°C 54°C 150°C Viscosity 6000cps 500cps 9000cps Colour clear/amber white brown Solvent Acetone or MEK Hot water 590-S or Methanol Wafer-Mount™ Product Wafer-Mount™ 559 Wafer-Mount™ 562 Description Semi-rigid solvent resistant plastic film with pressure sensitive soluble. Ideal for scribing wafers with vacuum hold down. Thermoplastic film adhesive with good adhesion to metals, glass, and ceramic and excellent for mounting fragile thin substrates. Form Sheet Sheet Size 0.005"x10"x10" 0.003"x8"x10" Weight n/a n/a Flow Point n/a 93°C Viscosity n/a n/a Colour clear white Solvent Acetone or MEK Trichloroethylene or Toluene APPLICATION PROCEDURES: Crystalbond™ 509, 555, 590 Heat a ceramic or glass mounting block to the flow temperature of the selected Crystalbond™ adhesive using a laboratory hot plate. The flow temperature for 509 is 77°C, 555-120°F, 590-320°F. Refer to Technical Bulletin M1 for information about Aremco's unfired ceramic mounting block. Aremcolx 502-1100. This ceramic is ground, flat and parallel, and ideal for dicing applications since the block tends to 'dress' or prepare the blade. Apply a uniform layer of the adhesive by pressing it on the heated backup block. Make sure the Crystalbond™ flow temperature is not exceeded, otherwise a degradation of the adhesive properties will result. Position the substrate on the backup block, allowing the substrate to heat up to the temperature of the block. Apply a small weight to ensure that the substrate settles evenly and that air bubbles are not drawn back under the substrate. Apply more Crystalbond™ around the edges of the substrate to create a fillet and increase strength. Remove the block/substrate assembly from the hot plate and allow it to cool slowly until the Crystalbond™ is hard. The weight can then be removed and the assembly cooled quickly to room temperature. Machine or process the substrate as required, then remove the parts by reheating the block to the Crystalbond™ flow temperature. Use a tool to slide the substrate or parts off the backup block. Since the block is now hot, the next substrate can be mounted. Clean the parts using the appropriate solvent in a three tank cleaning system: a dissolving tank to remove the bulk of the Crystalbond™, a wash tank to remove additional residue, and a final rinse tank. Ultrasonic cleaning may be used to facilitate cleaning. Conserve solvent by transferring from clean to contaminated tanks. When using 590-S stripper, mix 6-8oz. of stripper per gallon of water. Heat to 160 degrees F. Immerse parts a minimum of 5 minutes until Crystalbond™ 590 dissolves. Rinse in clear water. Safety Note: 590-S is caustic. Use eye protection and in the event of exposure , flush area immediately with water and see physician. WAFER-MOUNT™ 559: Peel clear plastic from backing paper. Position the substrate, face down, on the backing paper. Place the plastic film with the adhesive side down, over the part. Press firmly to assure good adhesion, then peel the backing paper away. After processing, remove the parts from the Wafer-Mount™ by heating at 300°F for 2-3 minutes until the adhesive softens. Refer to Crystalbond™ step 6 for cleaning instructions. WAFER-MOUNT™ 562: Cut the Wafer-Mount™ film adhesive to the required preform. Heat the backup block on the hot plate to 195-210 degrees F and melt the adhesive. Use multiple preforms if required to fill gaps created by warped substrates. Press the substrate down firmly and remove the heat. Once the adhesive is set, the assembly can be cooled in air or by water quenching, but care must be taken to avoid thermal shock cracking of the substrate. After processing, remove the parts from the Wafer-Mount™ by reheating the backup block to the flow temperature. Refer to Crystalbond™ step 6 for cleaning instructions. GENERAL SAFETY NOTE: Melting of Crystalbond™ and Wafer-Mount™ adhesives should be performed in a well ventilated area or fume hood.
DAPI-Fluoromount-G™
EMS17984-24 Dapi-Fluoromount-G™ is a water-soluble, instant-blue nuclear probe fluorescing (455nm) compound for mounting fixed slides using a staining procedure having an aqueous final step. Mounting slides with Dapi-Fluoromount-G™ stains the cell nucleus and may also reduce fluorochrome quenching during analysis of slides by fluorescence microscopy. This mounting medium also provides a semi permanent seal for long-term storage of slide preparations. Research Applications Slide Mounting Instructions For Use Prepare cytocentrifuge preps according to established procedures. Fix and rehydrate preps. Remove one slide at a time from aqueous buffer, add 1 drop of Dapi-Fluoromount-G™ directly to cell prep, mount cover slip and press gently with a gauze sponge to remove excess mounting medium and to seal the cover slip. Allow mounted preps to air-dry for 5 minutes before examination. Handling And Storage Dapi-Fluoromount-G™ is supplied as a 20 ml solution containing 0.1% NaN3 as preservative; store at room temperature. Avoid exposure to light. WARNING Reagents contain sodium azide which is very toxic if ingested or inhaled. Avoid contact with skin, eyes or clothing. Wear eye or face protection when handling. If skin or eye contact occurs, wash with copious amounts of water. If ingested or inhaled, contact a physician immediately. Sodium azide yields toxic hydrazoic acid under acidic conditions. Dilute azide-containing compounds in running water before discarding to avoid accumulation of potential explosive deposits in lead or copper plumbing. For Research Use Only. Not For Diagnostic Or Therapeutic Use.
DER 332-732 Epoxy Resin Embedding Kit
Introduction This kit combines two epoxy resins, DER 332, a very pure and uniform epoxy having a low viscosity, and DER 732, a polyglycol diepoxide which imparts softness and flexibility to the polymers of conventional epoxides, but does not cause shrinkage or serious loss of strength. Together with their curing agents, this kit produces a standard amber-coloured block which can be trimmed and sectioned easily. Recommended Procedure Fixation: Tissues can be fixed in a wide range of fixatives. One of the more commonly used fixatives is an aldehyde (i.e.: glutaraldehyde) followed by osmium tetroxide. Dehydration: There are many different dehydration schedules that can be followed. A typical one is as follows: 70% Ethanol for 10 minutes 100% Ethanol for 10 minutes 100% Ethanol for 15 minutes 100% Propylene Oxide for 15 minutes 100% Propylene Oxide for 15 minutes **NOTE: Longer times may be required for some samples. Mixing Instructions: Components Softer Harder Tougher, Soft A/E Ratio* DER 332 7 ml 7 ml 6 ml 7 ml DER 732 3 ml 2 ml 3 ml 3.2 ml DDSA 5 ml 5 ml 10 ml 8 ml DMP 30 0.3 ml 0.28 ml 0.38 ml 0.3 ml (FOR LARGER BATCHES INCREASE EACH COMPONENT PROPORTIONALLY) Slight variations of the accelerator (DMP-30) will drastically affect the colour and brittleness of the block. Prior to measuring and mixing, the resin and the anhydride should be warmed (60°C) to reduce their viscosity. Thorough mixing is imperative to be able to achieve uniform blocks. Although the mixture can be stored for up to 6 months at 4°C it is highly recommended that freshly prepared embedding medium always be used. If you choose to store the mixture you should warm it thoroughly prior to adding the accelerator. Infiltration: It is recommended that for all of the infiltration steps a specimen rotator be used. Drain the tissue of most of the propylene oxide, leaving a little so the tissue does not dry out. Replace the solvent with a 1:1 solution of propylene oxide:embedding medium and allow it to stand for at least 1 hour at room temperature. Remove the mixture, replace it with 100% embedding medium and leave for 6-12 hours at room temperature. Embedding: This may be done in EMS embedding capsules EMS70020 or a flat embedding mould (EMS70900). Transfer each sample to a dry capsule or mould and fill the mould with embedding medium. Curing of the medium may be carried out in an oven at 37°C, 45°C, and 60°C for 24 hours at each temperature. Blocks can be trimmed and sectioned after the blocks return to room temperature. ---------------------------------------------------------------------------------------------------------- *This formula was suggested by Bluemink (1970) in accordance with the recommendations of Coulter(1967) concerning anhydride/epoxy ratios. References Lockwood, W.R.(1964), Anatomical Record, 150, 129. Bluemink, J.G.(1970), J. Ultrastruct. Res. 32, 142. Product Information DER 332-732 Embedding Kits EMS embedding capsules Catalog #70020 Flat embedding mold Catalog #70900
DOW CORNING® High-Vacuum Grease, Silicone Compound
APPLICATIONS Seals and lubricates chemical processing equipment. Lubricating plug valves, control valves, flow meter bearings, ceramic plug cocks, fire extinguisher valves, water treating equipment, synthetic rubber gaskets and seals in high temperature applications. Sealing vacuum and pressure systems. Lubricating O-rings in binoculars and telescopes. Prevents fogging of delicate lenses. FEATURES Good lubricating and sealing ability Low volatility Excellent resistance to water, chemicals, high and low temperatures COMPOSITION Silicone oil Inorganic thickener TYPICAL PROPERTIES Specification writers: These values are not intended for use in preparing specifications. Please contact your local Dow Corning sales representative prior to writing specifications on this product. Standard* Test Result Colour White to grey, translucent Physical nature Stiff compound ISO 2137 Penetration, unworked 175 to 210mm/10 ISO 2137 Penetration, worked 60, max. 260mm/10 CTM 0033A A Bleed, 24 hours; 200°C <0.5% CTM 0033A B Evaporation, 24 hours; 200°C <0.2% Melting point None (°C) * CTM: Corporate Test Method, copies of CTMs are available on request. PERFORMANCE DATA Solubility DOW CORNING High-Vacuum Grease is insoluble in water, methanol, ethanol, acetone, glycol and glycerine. It can be dispersed in kerosene, stoddard solvent, benzene, toluene, ethyl ether or petroleum ether. Chemical resistance DOW CORNING High-Vacuum Grease is unaffected by most vegetable and minerals oils, many organic compounds, and most common gases. It is also resistant to most aqueous solutions of inorganic salts and to dilute acids and alkalies. The suitability of DOW CORNING High-Vacuum Grease should always be tested before the material is adopted for regular use. HANDLING PRECAUTIONS In case of eye contact, flush eyes with water. PRODUCT SAFETY INFORMATION REQUIRED FOR SAFE USE IS NOT INCLUDED. BEFORE HANDLING, READ PRODUCT AND SAFETY DATA SHEETS AND CONTAINER LABELS FOR SAFE USE, PHYSICAL AND HEALTH HAZARD INFORMATION. THE SAFETY DATA SHEET IS AVAILABLE FROM YOUR LOCAL DOW CORNING SALES REPRESENTATIVE. USABLE LIFE AND STORAGE When stored at or below 20°C in the original unopened containers, this product has a usable life of 60 months from the date of production. PACKAGING This product is available in different standard container sizes. Detailed container size information should be obtained from your nearest Dow Corning sales office or Dow Corning distributor. LIMITATIONS This product is neither tested nor represented as suitable for medical or pharmaceutical uses. HEALTH AND ENVIRONMENTAL INFORMATION To support customers in their product safety needs, Dow Corning has an extensive Product Stewardship organisation and a team of Health, Environment and Regulatory Affairs specialists available in each area. For further information, please consult your local Dow Corning representative. WARRANTY INFORMATION - PLEASE READ CAREFULLY The information contained herein is offered in good faith and is believed to be accurate. However, because conditions and methods of use of our products are beyond our control, this information should not be used in substitution for customer's tests to ensure that Dow Corning's products are safe, effective, and fully satisfactory for the intended end use. Dow Corning's sole warranty is that the product will meet the Dow Corning sales specifications in effect at the time of shipment. Your exclusive remedy for breach of such warranty is limited to refund of purchase price or replacement of any product shown to be other than as warranted. Dow Corning specifically disclaims any other express or implied warranty of fitness for a particular purpose or merchantability. Unless Dow Corning provides you with a specific, duly signed endorsement of fitness for use, Dow Corning disclaims liability for any incidental or consequential damages. Suggestions of use shall not be taken as inducements to infringe any patent. ©1997-2001 Dow Corning Corporation. All rights reserved. 23/11/1998. Ref. no. 22-0911C-01 DC 3677 ® DOW CORNING is a registered trademark of Dow Corning Corporation.
DPX Mounting Medium
A mixture of distyrene (a polystyrene), a plasticiser (tricresyl phosphate), and xylene, called DPX, was introduced in 1939 and later modified by the substitution of a more satisfactory plasticiser, dibutylphthalate (butyl, phthalate, styrene - BPS) This colourless, synthetic resin mounting media is now available at ProSciTech and has generally replaced xylene-balsam. It preserves stains and dries quickly. Surplus mountant may be peeled off the preparation after cutting around the coverslip with a razor blade or a scalpel. It is not recommended for use with thick sections (e.g. cellulose nitrate) where there is a danger of retraction of the mountant upon drying. For these Canada Balsam (IA158) is a suitable alternative. ENTELLAN® NEW (IM022) replaces DPX (Depex).
Durcupan™ ACM Epoxy Resin
Araldite Base Embedding Agent for Electron Microscopy Araldite casting resin M, hardener grade 964, accelerator grade 964, and dibutyl phthalate, the components required for the Araldite embedding procedures as described in the Literature are sold by us under the description Durcupan ACM. Mixtures of Durcupan ACM present an interesting alternative to methacrylates for embedding sections for electron microscopy. They present the advantage that they harden uniformly, practically without shrinkage. All fixing agents normally used for electron microscopic work are also suitable for the Durcupan ACM embedding process. This process consists of three phases: dehydration of the tissue with acetone or ethyl alcohol (1), infiltration with the embedding agent, followed by hardening. The following two resin mixtures have been found suitable in practice: No. 1 Durcupan ACM mixture: Proportion in the mixture Component Identification color (bottle cap and label) Product 10ml A/M Red Epoxy resin 10ml B Blue 964 hardener 0.1-0.2ml D Green Dibutyl phthalate (2) No. 2 Durcupan ACM mixture: 10ml A/M Red Epoxy resin 10ml B Blue 964 hardener 0.3-0.4ml C Yellow 964 accelerator 0.1-0.2ml D Green Dibutyl phthalate (2) These two mixtures are preferably prepared at least 15 minutes before use, and kept during this period in the drying cupboard at 50°C, so that they are thoroughly mixed. Glass apparatus used for measuring and mixing should be placed in acetone or absolute ethyl alcohol immediately after use and cleaned. (1) Durcupan ACM mixtures are not as easily soluble in ethyl alcohol as in acetone. (2) This reduces the brittleness of the blocks and improves their cutting properties. Basic scheme for dehydration and embedding in Durcupan ACM The tissues are dehydrated, after they have been fixed in buffered osmic acid (osmium tetroxide) or in any other fixing agent normally used for electron microscopy. Normal dehydration stage: No. 1 tray 30% acetone (3) 15 minutes With materials containing only very little water it is permissible to start directly with No. 2 tray No. 2 tray 50% acetone (3) 30 minutes No. 3 tray 70% acetone (3,4) 30 minutes This period may also be extended, e.g., during lunch time or overnight. No. 4 tray 90% acetone (3) 30 minutes No. 5 tray Dry acetone (3) 30 minutes Dry acetone is stored over dried copper sulphate. No. 6 tray Dry acetone (3) 30 minutes Normal infiltration stage: No. 7 tray 3 parts dry acetone (3) 1 part of No. 1 Durcupan ACM mixture 1 hour @ room temperature No. 8 tray 2 parts of dry acetone(3) 2 parts of No. 1 Durcupan ACM mixture 1 hour @ room temperature No. 9 tray 1 part dry acetone (3) 3 parts of No. 1 Durcupan ACM mixture 1 hour @ room temperature (this period may also be extended, during lunch time or overnight) No.10 tray No. 1 Durcupan ACM mixture 1-2 hours @ 50°C in the drying cupboard No. 11 tray No. 1 Durcupan ACM mixture 1-2 hours @ 50°C in the drying cupboard No. 12 tray No. 2 Durcupan ACM 1-2 hours @ 50°C in the drying cupboard (3) Instead of acetone, ethyl alcohol of the same concentration may be used. (4) Aqueous solutions of the more common contrasting agents can be added to No. 3 tray. Normal hardening stage: Take the pieces of tissue from No 12 tray and place them in dry gelatin capsules, which are then filled with No 2 Durcupan ACM Mixture (from the storage vessel). After closing the capsules, harden them in the drying cupboard at 50-80°C for at least 48 hours. In this way, blocks of a pale golden colour are obtained, of hardness similar for that of methacrylate, with good cutting properties. If contrasting is required and this has not been carried out during dehydration (No. 3 tray), let the sections float on a heavy metal salt solution. Unlike methacrylate, hardened Durcupan ACM cannot be dissolved out of the sections by applying gentler media. For details as regards dehydration, embedding, hardening and cutting refer to the book of Daniel C. Pease, Histological Techniques for Electron Microscopy, Academic Press Inc., New York and London, 1960 (pp. 78-85). Caution Take great care when working with Durcupan ACM: Do not breathe in the vapor and avoid skin contact, because this may cause skin irritation and allergic reactions. Splashes on the skin must be washed off immediately with a 3% boric acid solution. Frequent washing of hands, arms, and face with lukewarm soap water is advisable.
EMS Glycerol Mounting Medium
EMS17989-40, EMS17989-41 This mounting medium is made with glycerol. It is excellent for preserving thick sections that can not be mounted with mounting medium or fresh frozen tissues that contain a lot of lipids, e.g. brain tissue. It is also useful for preserving small embryos. This mounting medium is also useful for immunofluorescence. This is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy dyes, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine and Redox. The fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes. Not recommended for phycoethyrin (PE), phycocyanin (PC) and allophycocyanin (APC). Intended Use Mounting tissues and cell for IHC and for Immunofluorescence (IF) Reagent Ready to use mounting medium for IHC and for Immunofluorescence (IF) Refractive Index 1.472 Storage 2-8°C is recommended, Protect from light Procedure Dehydration of specimen is not required. Bring the vial to room temperature. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 1-3 drops of mounting medium directly on top of the specimen. Place coverslip; it may be necessary to seal the edges with nail polish or any organic mounting medium. Method for applying Coverslip: Put 1-2 drops of Mounting medium on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. For storage it is recommended that the IHC slides can be stored at room temperature and IF slides in the dark at 2-8°C.
EMS Glycerol Mounting Medium with DABCO™
EMS17989-50, 17989-51 Glycerol with antifading agent 1, 4-Diazobicyclo-(2,2,2-octane (DABCO™) is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Fluorescence is retained during prolonged storage at 4°C in the dark. Not recommended for phycoethyrin (PE), phycocyanin (PC) and allophycocyanin (APC). Intended Use Immunofluorescence, confocal microscopy Reagent Ready to use mounting medium with anti-fading agent Refractive Index 1.4615 Storage 2-8°C is recommended, Protect from light Procedure Dehydration of specimen is not required. Bring the vial to room temperature. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 1-2 drops of mounting medium directly; cover slip carefully avoiding air bubbles. The specimen is ready for visualization under a microscope. One can seal the edges of cover slip with nail polish or any organic medium. Method for applying Coverslip: Put 1-2 drops of Mounting medium on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. For storage it is recommended that the slide be stored in the dark at 2-8°C.
EMS Glycerol Mounting Medium with DAPI and DABCO™
EMS17989-60, EMS17989-61 Glycerol with DAPI and DABCO™ is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Fluorescence is retained during prolonged storage at 4°C in the dark. This mounting medium is fortified with DAPI which is a counter-stain for DNA. This product is to be used in situ hybridization techniques or other methods where fluorescence of DNA staining is required. DAPI excites at 360nm and emits at 460nm, producing a blue fluorescence. RNA is also stained with DAPI. Glycerol with antifading agent 1, 4-Diazobicyclo-(2,2,2-octane (DABCO™) is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of fluorochromes. The fluorescence is retained during prolonged storage at 4°C in the dark. Not recommended for phycoethyrin (PE), phycocyanin (PC) and allophycocyanin (APC). Intended Use Immunofluorescence, confocal microscopy Reagent Ready to use mounting medium with anti-fading agent Refractive Index 1.4617 Storage 2-8°C is recommended, Protect from light Procedure Dehydration of specimen is not required. Bring the vial to room temperature. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 1-2 drops of mounting medium directly; cover slip carefully avoiding air bubbles. The specimen is ready for visualization under a microscope. One can seal the edges of cover slip with nail polish or any organic medium. Method for applying Coverslip: Put 1-2 drops of Mounting medium on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. For storage it is recommended that the slide be stored in the dark at 2-8°C.
EMS Mounting Medium with DAPI and PG
EMS17989-30, EMS17989-31 EMS Mounting Medium With 4,6-diamidino-2-phenylindole (DAPI) and Propyl Gallate (PG) Fluoroshield with DAPI is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Phycoerythrin (RP-E), Phycocyanin (PC), and Allophycocyanin (APC). Fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP-E, PC and APC. This mounting medium is fortified with DAPI which is a counter-stain for DNA. This product is to be used in situ hybridisation techniques or other methods where fluorescence of DNA staining is required. DAPI excites at 360nm and emits at 460nm, producing a blue fluorescence. RNA is also stained with DAPI. May encounter problems with frozen brain or other frozen tissues with lots of fat. Intended Use Immunofluorescence, confocal microscopy Reagent Ready to use mounting medium Refractive Index 1.364 ± 0.002 (This number applies to this mounting medium in solution. Refractive indexes change when the water solvent evaporates and mounting media dries on slides. We do not have the means to measure the refractive indexes of dry mounting mediums; however, we expect the numbers to go higher when dried. The refractive index of water is 1.3330.) Storage 2-8°C, Protect from light, DO NOT FREEZE Procedure Bring the vial to room temperature. Rinse slide to be mounted with distilled or deionised water; touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium directly on top of the specimen. Let stand at room temperature for about 5 minutes in the dark. Apply cover slip, carefully avoiding air bubbles. The specimen is ready for visualisation under a microscope. One can seal the edges of the cover slip with nail polish, or any organic mounting medium. If a coverslip is not sealed, air bubbles will appear in few days. Method for applying Coverslip: Put 1-2 drops of FLS on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. Incubate at 37°C for one hour in the dark to dry organic mounting medium. For long term storage it is recommended that the slide be stored in the dark at 2-8°C. Removal of Coverslip: Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) water for a few minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
EMS Shield Mounting Medium with DAPI and DABCO™
Fluoroshield with DAPI is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Phycoerythrin (RP-E), Phycocyanin (PC), and Allophycocyanin (APC). Fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP-E, PC and APC. This mounting medium is fortified with DAPI which is a counter-stain for DNA. This product is to be used in situ hybridisation techniques or other methods where fluorescence of DNA staining is required. DAPI excites at 360nm and emits at 460nm, producing a blue fluorescence. RNA is also stained with DAPI. You may encounter problems with frozen brain or other frozen tissues with lots of fat. Intended Use Immunofluorescence, confocal microscopy Reagent Ready to use mounting medium Refractive Index 1.364 ± 0.002 (This number applies to this mounting medium in solution. Refractive indexes change when the water solvent evaporates and the mounting media dries on slides. We do not have the means to measure the refractive indexes of dry mounting mediums; however, we expect the numbers to go higher when dried. The refractive index of water is 1.3330.) Storage 2-8°C, Protect from light, DO NOT FREEZE Procedure Bring the vial to room temperature. Rinse the slide to be mounted with distilled or deionized water; touch the edges of the slide with a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium directly on top of the specimen. Let it stand at room temperature for about 3-5 minutes in the dark. Apply the cover slip, carefully avoiding air bubbles. The specimen is ready for visualization under a microscope. One can seal the edges of the cover slip with nail polish or any organic mounting medium. If a coverslip is not sealed, air bubbles will appear in few days. Method for applying Coverslip: Put 1-2 drops of FLS on the specimen. After 3-4 minutes apply the coverslip carefully, avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. Incubate at 37°C for one hour in the dark to dry organic mounting medium. For long term storage, it is recommended that the slide be stored in the dark at 2-8°C. Removal of Coverslip: Coverslip can be removed before sealing the edges. Soak the slide in warm (37°C) water for a few minutes. Carefully and slowly, move the coverslip. Soak in water for an additional few minutes to remove the coverslip. Rinse the slide several times with warm water to remove all the mounting medium. The slide can be remounted again.
EMS Shield Mounting Medium with DAPI and PG
EMS17989-30, 17989-31 Fluoroshield with DAPI is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Phycoerythrin (RP-E), Phycocyanin (PC), and Allophycocyanin (APC). Fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP-E, PC and APC. This mounting medium is fortified with DAPI which is a counter-stain for DNA. This product is to be used in situ hybridisation techniques or other methods where fluorescence of DNA staining is required. DAPI excites at 360nm and emits at 460nm, producing a blue fluorescence. RNA is also stained with DAPI. May encounter problems with frozen brain or other frozen tissues with lots of fat. Intended Use Immunofluorescence, confocal microscopy Reagent Ready to use mounting medium Refractive Index 1.364 ± 0.002 (This number applies to this mounting medium in solution. Refractive indexes change when the water solvent evaporates and mounting media dries on slides. We do not have the means to measure the refractive indexes of dry mounting mediums; however, we expect the numbers to go higher when dried. The refractive index of water is 1.3330.) Storage 2-8°C, Protect from light, DO NOT FREEZE Procedure Bring the vial to room temperature. Rinse slide to be mounted with distilled or deionized water; touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium directly on top of the specimen. Let stand at room temperature for about 5 minutes in the dark. Apply cover slip, carefully avoiding air bubbles. The specimen is ready for visualisation under a microscope. One can seal the edges of the cover slip with nail polish, or any organic mounting medium. If a coverslip is not sealed, air bubbles will appear in few days. Method for applying Coverslip: Put 1-2 drops of FLS on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. Incubate at 37°C for one hour in the dark to dry organic mounting medium. For long term storage it is recommended that the slide be stored in the dark at 2-8°C. Removal of Coverslip: Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) water for a few minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
EMS-Mount Mounting Medium
EMS17985-05/-06/-07/-08 EMS Mount Mounting Medium is a non-permanent, low-viscosity aqueous mounting medium designed for mounting of tissue sections and cell smears with peroxidase and alkaline phosphatase chromogens that cannot be dehydrated with organic solvents. This mounting medium preserves Fast red, Aminoethylycarbazole (AEC), BCIP/NBT, BCIP/INT chromogens and is also compatible with counterstain like Haematoxylin and Nuclear fast red (NFR). It is also suitable for chromogens like DAB and DAB with nickel and cobalt. (It is not compatible with H and E staining). Coverslip is required. Refractive Index 1.400 ± 0.0021 (1These numbers apply to these mounting mediums in solution. Refractive indexes change when the water solvent evaporates and mounting media dries on slides. We do not have the means to measure the refractive indexes of dry mounting mediums; however we expect the numbers to go higher when dried. The refractive index of water is 1.3330 Applications Mounting of chromogens that cannot be dehydrated with organic solvents. Reagent Ready to use mounting medium Storage Storage at 2-8°C is recommended. DO NOT FREEZE. Instructions For Use Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONIZED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium and apply coverslip carefully without getting air bubbles. The specimen is ready for visualization under a microscope. The coverslip may move, so it may be necessary to seal the edges of cover slip with nail polish, any organic medium or our Organo mounting medium Method for applying Coverslip: Put 1-2 drops of Mounting medium on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium on the edges to seal. Incubate at 37°C for one hour in the dark to dry organic mounting medium. Store slide at room temperature. Removal of Coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) distilled or deiononized water for several minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
Entellan New Mounting Medium (DG)
Entellan® New is a solution of polymers in xylene. Application: The combination of several synthetic resins makes Entellan® a particularly rapid embedding agent. It can be used for all dehydrated microscopic preparations. Embedded preparations can be filed for years without changes occurring in their color. Constant properties ensure its high quality. Properties: Setting time at room temperature 20 minutes Intrinsic colour Colourless; absolutely transparent Neutrality Acid number <2.50 Refractive Index nd20 1.490 - 1.500 Refraction to daylight & UV light not turbid Heat resistance up to 90°C Cold resistance down to -17°C Intrinsic fluorescence under UV light <50ppb quinine sulfate Indifference to biological stains not to carmine, sudan III , toluidine Density 20 o/4°C 0.940 - 0.960 Viscosity @ 20°C <450cP Solubility dissolves in acetone, ether, benzene, chloroform, dioxan, toluene, xylene Reaction to water slight; immiscible with water Loss on drying about 75% Use: Cover the object with drops of Entellan® so that the space between the cover glass and the slide is completely filled. The cover glass must be sealed right up to each edge. Excess embedding agent can be removed with a cloth moistened with xylene. In the case of histological and cytological preparations, it is recommended that xylene be used as the final stage before embedding, so that turbidities caused by the action of alcohol can be avoided. As a matter of precaution, we recommend that, as with other embedding agents, all work with Entellan® be carried out in a fume hood or in a well-ventilated room. Note: Recap the bottle immediately after use. Should Entellan® become to viscous, it can be brought back to the desired consistency by adding xylene. Store at room temperature
Fluka Durcupan
Fluka Durcupan, the new embedding agent, opens new roads in the observation of enzymatic digestion processes and for carrying out histochemical investigations in the submicroscopic range. It enables formalin and other aqueous fixatives to be used for electron microscopy and the dehydration of the tissue in alcohol or acetone to be dispensed with, so that excessive changes in the ultrastructure of the cells are obviated. Fluka Durcupan is supplied in original packages, each package containing enough material for 10 embedding operations. The material comprises the following four components: Component Contents A 100g of embedding substance, a water-soluble aliphatic polyepoxide (the former Experimental Product X 133/2097 of the CIBA Aktiengesellschaft, Base, Switzerland) B 100g of 964 hardener, an anhydride of a diazide with aliphatic side chain C 20g of 960 hardener, a phenol derivative with amino groups D 20g of plasticizer (dibutyl phthalate) These four components make up a polymerisable mixture. Directions 1. Fixing and dehydrating Fix the tissue in, say, osmium tetroxide, potassium permanganate, or 10% formaldehyde (with Veronal buffer, pH 7 - 7.5). Dehydrate it not in alcohol or acetone, but in a series of mixtures of water with Component A, the concentration of Component A increasing in consecutive stages, as shown in the following table: Dehydration Bath 1st Tray 50% Component A with 50% water* for 30 minutes shake tray if possible 2nd Tray 70% Component A with 30% water* for 45 minutes** shake tray if possible 3rd Tray 90% Component A with 10% water* for 45 minutes** shake tray if possible 4th Tray 100% Component A for 90 minutes** shake tray if possible 5th Tray 100% Component A for 90 minutes** shake tray if possible * Aqueous solutions of the following contrasting agents may be added to the contents of these trays: Osmium tetroxide, phosphotungstic acid, uranyl acetate. Do not add lead hydroxide (see section on Contrasting). ** These times may be varied without danger (e.g. up to 90 minutes in 2nd tray). 2. Embedding The dehydrated tissue is now placed in a polymerisation mixture of the following composition: Proportion Component Miscibility with Water 5ml A all proportions 11.7ml B nil 1.0 - 1.2ml (very critical!)*** C limited 0.2 - 0.4ml D nil *** Insufficient Component C: The block becomes too soft. Excess Component C: The block becomes granular Leave the tissue overnight at 4 - 5°C in this comparatively viscous mixture. Its is advisable to shake it while in the mixture. 3. Polymerisation Now fill the gelatine capsules with the freshly prepared polymerisation mixture of the above composition and charge them with the tissue specimens. Polymerisation takes place at 37 - 45°C and lasts 3-4 days. Contrasting The tissue fixed with formaldehyde shows a very low contrast in the electron microscope, an advantage for certain histochemical investigations. However, for purely morphological investigations, contrasting agents must be used. The following heavy-metal compounds are suitable for this purpose: osmium tetroxide, phosphotungstic acid, lead hydroxide and uranyl acetate. Contrasting can be carried out in two ways: Either let the mount mesh covered with ultrathin specimens float for about an hour on solutions of these heavy-metal compounds, or add the aqueous heavy-metal compound solution (but not lead hydroxide) to the dehydration bath trays marked \*. The blocks obtained according to this embedding method with Fluka Durcupan are mostly softer than methacrylate blocks and present certain difficulties in the preparation of ultra-thin sections, particularly because of the chatter, a frequent occurrence. Ordinary glass knives or knives made of the harder Tempax glass are suitable for ultramicrotomy. (Diamond knives have not produced any better results than glass knives.) The sections can be caught in a solution of 20 - 50% acetone in water. If the Durcupan blocks are too soft, try to cut the tissue out and rebed it in prepolymerized methacrylate. Such Durcupan / methacrylate blocks are usually much easier to cut. Animal tissue can be embedded in Fluka Durcupan more easily than plant tissue. Caution Take great care when working with Durcupan, as this substance may cause skin irritation and allergic reactions. Work always with rubber gloves. Immediately wash off droplets on the skin with 3% boric acid solution. Wash hands, arms, face frequently in lukewarm soap water.
Fluoro-Gel II with DAPI Mounting Medium
EMS17985-50, EMS17985-51 Description Fluoro-Gel II is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox, phycoerythrin (R-PE), phyocyanin (PC), and allophycocyanin (APC). The fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, R-PE, PC and APC. Applications Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium. Instructions For Use Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 3-4 drops of mounting medium directly on top of the specimen and spread out evenly by tilting slide back and forth or spread evenly with a 0.2 ml plastic pipette tip making sure the tissue is not touched. Excess medium can be removed by touching the edges of slide against paper towel. Let stand at room temperature for about 5 minutes. Apply cover slip carefully avoiding air bubbles. One can seal the edges of cover slip with nail polish, any organic medium or our Limonene mounting medium. If a coverslip is not sealed air bubbles will appear in few days. For storage it is recommended that the slide be stored in the dark at 2-8°C. Handling And Storage This product can be store at room temp, however for long term storage 2-8°C is recommended. Protect from light, DO NOT FREEZE Removal of Coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) distilled or deionised water for several minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
Fluoro-Gel III
Description Fluoroshield mounting medium with PI is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa Fluoro 594, Green Fluorescent protein (GFP), tetramethyl rhodamine, Redox, Phycoerythrin (RP‐E), Phyocyanin (PC), and Allophycocyanin (APC). The fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP‐E, PC and APC. This mounting medium is fortified with PI, which is a counter‐stain for DNA. This product is to be used in in situ hybridisation techniques or other methods where fluorescence of DNA staining is required. PI excites at 535nm and emits at 615nm, producing a red fluorescence. RNA is also stained with PI. Intended use Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium. Storage 2‐8°C, protect from light, DO NOT FREEZE. Procedure Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED or DEIONIZED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and remove any air bubbles. Apply 3‐4 drops directly of mounting medium on top of the specimen and spread out evenly by tilting the slide back and forth or spread evenly with a 0.2mL plastic pipette tip, making sure the tissue is not touched. Excess medium can be removed by touching the edges of slide on a paper towel. Let stand at room temperature for about 5 minutes. Apply cover slip, carefully avoiding air bubbles. The specimen is ready for visualisation under microscope. One can seal the edges of cover slip with nail polish or any organic medium or Organo Mounting Medium. If coverslip is not used air bubbles will appear in a few days. 9. For long‐term storage it is recommended that the slide be stored in the dark at 2‐8°C. Removal of coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) water for a few minutes. Carefully and slowly move the coverslip. Soak in water an additional few minutes and remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted.
Fluoro-Gel with DABCO™
Description Fluoro‐Gel with antifading agent 1,4‐Diazobicyclo‐(2,2,2)‐octane (DABCO™) is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Alexa fluoro 488, Alexa fluoro 594, Cy dyes, tetramethyly rhodamine, and Redox. The fluorescence is retained during prolonged storage at 4°C in the dark. Intended use Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium with anti‐fading agent. Storage 2‐8°C is recommended. Protect from light, DO NOT FREEZE. Procedure Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 3‐4 drops of mounting medium directly on top of the specimen and spread out evenly by tilting slide back and forth or spread evenly with a 0.2 mL plastic pipette tip, making sure the tissue is not touched. Excess medium can be removed by touching the edges of slide against paper towel. Let stand at room temperature for about 5 minutes. Apply cover slip, carefully avoiding air bubbles. The specimen is ready for visualisation under a microscope. One can seal the edges of cover slip with nail polish, any organic medium or our Organo mounting medium. If a coverslip is not sealed air bubbles will appear in a few days. It is recommended that the slide be stored in the dark at 2‐8°C. Removal of coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) distilled or deionised water for several minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted. Note: For research use only; not for use in diagnostic procedures. FOR IN VITRO LABORATORY USE ONLY.
Fluoro-Gel with Para Phenylenediamine (PPD) Anti Fading Mounting Medium
IM037-100 Description: Fluoroshield with strong antifading agent, 1,4-phenylenediamine (PPD) is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Alexa fluoro 488, Alexa fluoro 594, tetramethyly rhodamine, and Redox. The fluorescence is retained during prolonged storage at 4°C in the dark. This medium contains phenylenediamine and is not suitable for immunofluorescence of Cy dyes, GFP, mCherry, Phycoerythrin (R-PE), phyocyanin (PC), and allophycocyanin (APC) and other fluorochromes proteins. Refractive Index 1.366 ± 0.002 (This number applies to this mounting medium in solution. Refractive indexes change when the water solvent evaporates and mounting media dries on slides. We do not have the means to measure the refractive indexes of dry mounting mediums; however, we expect the numbers to go higher when dried. The refractive index of water is 1.3330) May encounter problems with frozen brain or other frozen tissues with lots of fat. Intended Use Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium with dark coffee colour. This colour does not interfere with Immunofluorescence. Storage 2-8°C is recommended. Protect from light, DO NOT FREEZE. Procedure Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium directly on top of the specimen. Let stand at room temperature for about 3-5 minutes in the dark. Apply cover slip carefully avoiding air bubbles. The specimen is ready for visualisation under a microscope. One can seal the edges of cover slip with nail polish, any organic medium. If a coverslip is not sealed air bubbles will appear in few days. Method for applying Coverslip: Put 1-2 drops of Mounting medium on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. Incubate at 37°C for one hour in the dark to dry organic mounting medium. For storage it is recommended that the slide be stored in the dark at 2-8°C. Removal of Coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) distilled or deionised water for several minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slides several times with warm water to remove all mounting medium. The slide can be remounted again. Warning For research use only; not for use in diagnostic procedures. FOR IN VITRO LABORATORY USE ONLY “In vitro laboratory products for research”
Fluoro-Gel with Tris Buffer
Description Fluoro-Gel is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photo-bleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Phycoerythrin (RP-E), phyocyanin (PC), and allophycocyanin (APC). The fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP-E, PC and APC. Intended use Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium. Storage 2-8°C, protect from light, DO NOT FREEZE. Procedure Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 3-4 drops of mounting medium directly on top of the specimen and spread out evenly by tilting slide back and forth or spread evenly with a 0.2 mL plastic pipette tip making sure the tissue is not touched. Excess medium can be removed by touching the edges of slide against paper towel. Let stand at room temperature for about 5 minutes. Apply cover slip carefully avoiding air bubbles. The specimen is ready for visualisation under a microscope. One can seal the edges of cover slip with nail polish, any organic medium or EMS Limonene Mount (17987-01) medium. If a coverslip is not used, air bubbles will appear in few days. For long term storage it is recommended that the slide be stored in the dark at 2-8°C Removal of coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) water for a few minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted.
Fluoromount-G™
EMS17984-25 Fluoromount-G™ is a water-soluble, non-fluorescing compound for slides mounted after a staining procedure having an aqueous final step. Mounting slides with Fluoromount-G™ may reduce fluorochrome quenching during analysis of slides by fluorescence microscopy. This mounting medium also provides a semi-permanent seal for long-term storage of slide preparations. Applications Slide Mounting Instructions For Use Prepare cytocentrifuge preps according to established procedures. Fix and rehydrate preps. Remove one slide at a time from aqueous buffer, add one (1) drop of Fluoromount-G™ directly to cell prep, mount cover slip and press gently with a gauze sponge to remove excess mounting medium and to seal the cover slip. Allow mounted preps to air-dry for five (5) minutes before examination. Handling And Storage Fluoromount-GT is supplied as a 25 ml solution containing 0.1% sodium azide as preservative; store at room temperature. Removal of Fluoromount™ Fluoromount™ is an aqueous based mounting medium and can easily be removed by soaking the slides in deionised water: Place the slide in a beaker full of deionised water on a magnetic stirrer. Leave the slide for few hours to overnight with gentle stirring for complete removal of dry Fluoromount™ Warning Reagents contain sodium azide, which is very toxic if ingested or inhaled. Avoid contact with skin, eyes, or clothing. Wear eye or face protection when handling. If skin or eye contact occurs, wash with copious amounts of water. If ingested or inhaled, contact a physician immediately. Sodium azide yields toxic hydrazoic acid under acidic conditions. Dilute azide-containing compounds in running water before discarding to avoid accumulation of potentially explosive deposits in lead or copper plumbing.
Formvar Resin
Formvar Resin is soluble in ethylene dichloride, xylene or dioxane. It is difficult to maintain a given percentage of solution due to the constant evaporation of the solvent. It is recommended that only small amounts of solution be prepared at a time. For most purposes a 0.25% solution will suffice for most of the substrate requirements. Several points to keep in mind: When a drop is spread on water there is a variation of thickness from the centre out. When a slide is dipped in the solution the lower or drain end will tend to show some variation in thickness. When not in use solutions should be tightly capped. If moisture gets into the solution, holey films will result. A rule of thumb for various thickness films is that 0.1 grams of Formvar in 100ml of solvent will produce films 10nm thick, 0.2g - 20nm, 0.5g - 50nm etc. To make a 0.25% Formvar solution: Dissolve 0.25g Formvar in 100ml high grade ethylene dichloride. Store solution in a well-stoppered brown glass bottle.
Fro-Marker and Para-Marker Pen
EMS71313-10 & EMS71313-20 Usage of the Fro-Marker & the Para-Marker Features: These pens are designed to make the production of coherent high quality immunocytochemical preparations more effective, accurate and with much less waste. They have applications for frozen paraffin and cytological preparations. As well, they prevent frozen sections and paraffin sections from moving, wrinkling, or falling off of the slide during immunostaining procedures. Warning: Cap tightly when not in use!!! Processing: Each membrane, which is applied by the marker, should be allowed to dry completely( approximately 5 minutes) at room temperature prior to routine processing. Warning: To avoid getting background during immunostaining do not use the markers at temperatures lower than room temperature.
Glutaraldehyde - Carbohydrazide (GACH)
This water miscible (aqueous) embedding medium prevents the loss of lipids caused by organic solvents and also the clouding of tissue observed during specific reactions involving the use of metallic fixatives (e.g., OsO4 and Potassium Permanganate). The GACH embedding medium is prepared by adding Carbohydrazide to 50% Glutaraldehyde in a final concentration of 150 mg/ml, (e.g., 1.5g of Carbohydrazide to 10mL of 50% Glutaraldehyde). The mixture is carried out in a pre‐cooled beaker kept in an ice bath, using a magnetic stirrer. The Carbohydrazide is added in three portions, and stirred rapidly for 15 minutes after each of the additions. It can be employed at neutral pH and cured at 37°C. The stock solution can be stored for several months at ‐20°C without deterioration. Tissue Preparation: Fixation: Tissues are fixed in the conventional way with concentrations of 2 to 5% Glutaraldehyde and rinsed with the buffers. Dehydration: Dehydrate according to the following schedule with aqueous dilution's of GACH. 20% GACH 80% H20 ......... 3 hours or overnight 50% GACH 50% H20 ......... 2 ‐ 4 hours 80% GACH 20% H20 ......... 2 ‐ 4 hours 100% GACH ......... 1 ‐ 2 hours This procedure is carried out in a cold room or in vials immersed in ice (1°C). Embedding: Particular specimens, (e.g., erythrocytes) can be embedded in the undiluted medium, suspended directly in 100% GACH, [Dodge, et al]). Embedding is carried out by removing the tissue from the last 100% GACH and placing it on a small droplet of fresh GACH on a dental wax plate at room temperature. Polymerisation is obtained by placing the plate in an incubator at 37°C for 8 ‐ 14 hours. Sectioning: The droplets of polymerised GACH can be cemented on blank epoxy blocks, trimmed and then sectioned on the ultramicrotome using diamond or glass knives. Polymerised GACH is no longer soluble in water; therefore, the sections can float on the surface of the water in the boat of the knife. The thickness of the sections can be judged by interference colour similar to epoxy sections. To increase contrast on the cellular components, sections can be stained with aqueous Uranyl Acetate and Lead Citrate. Reference: Dodge, J. T.; Mitchel, C. ‐ Arch. Biochem. biophys. 100 ‐ 119 (1963).
Glycol Methacrylate (GMA) Embedding Medium
Glycol Methacrylate (GMA) was introduced as an embedding medium for ultrastructural cytochemical studies in Electron Microscopy. An improved method for GMA embedding was described by Leduc & Bernhard(1967). This method provides a better preservation of the tissues and was more useful for enzymatic extraction and autoradiographic studies. The following is the procedure which was recommended by Leduc & Bernhard. RECOMMENDED PROCEDURE: Fixation: Tissue should be fixed in an 1.25% Glutaraldehyde in 0.1M Sodium Cacodylate or Phosphate buffer, pH 7.2, for 1 hour. The tissue should be rinsed in the same buffer for 1 hour or overnight. Dehydration and Infiltration: 80% of GMA monomer and 20% distilled water for 15 minutes 100% GMA- 4 changes at 15 minutes each change Embedding medium catalyst for 1 hour. Embedding medium catalyst for 1 hour. Final infiltration in partially polymerised embedding medium overnight. NOTE: All fixation and infiltration, embedding and final polymerisation with UV light are carried out in a cold room at 3ºC or on top of ice. Mixing Instructions: Mixture of 97% GMA plus 3% distilled water Mixture of 98% butyl methacrylate plus 2% 2,4 dichlorobenzoyl peroxide (Luperco) Final Mixture: 7 parts or 70ml of mixture 1 3 parts or 29.4ml BMA of Mixture 2 and 0.6g Luperco To reduce the swelling of artifacts, the above mixture is partially polymerised before use. It should be of the consistency of maple syrup. The prepolymer is prepared as follows: Place a small amount of the above mixture in a large, capped flask, while heating over a bunsen burner with very rapid swirling until it boils. This should take approximately 1 minute. The flask is plunged immediately into a bath of ice water and agitated vigorously until it cools to about 2°C. If the initial viscosity is lower than that of the consistency of maple syrup, the heating and cooling process should be repeated for several times. The entire process takes about 5 minutes and the prepolymer may be stored in the freezer indefinitely. Embedding: The tissue is placed in gelatin capsules (NOT POLYETHYLENE), filled to the top with fresh prepolymer. Capsules should be closed, leaving as little air as possible. Capsules should be held upright in supports which permit the maximum passage of UV light. With long wave UV light, polymerisation takes from 25-48 hours, depending upon the viscosity of the prepolymer, the amount of accelerator added, and the source of UV light. Sections should be picked up only on coated grids which can be stained with uranyl acetate or lead acetate. Tissues embedded in GMA have very dense, nucleic acid containing structures. GMA is also useful as an embedding medium for sectioning of tissue for light microscopy(1-2 microns). REFERENCES: Leduc, E. & Bernhard, W.(1967), Ultrastructure Research 4, 196-199. Rosenberg, M., Bartl, P., and Lesko, JR.(1960), Ultrastructure Research 4, 298.
Gold Conjugates
Gold Conjugates, a superior gold probe for light and electron microscopy. Probe Stability - Long term stability when stored at -25°C or less. Clustering - 85% singlets with no clusters greater than triplets Concentration and Dilution - EM and LM probes can be diluted from 1:10 to 1:200. Blotting probes can be diluted from 1:100 to 1:500 Size Distribution - EM grade probes have a low coefficient of variation, making them quite suitable for multiple labelling applications Particle Sizes - Unconjugated colloidal gold sols come in a wide range of sizes: 2, 5, 10, 15, 20, 30, 40, 50, 60, 80, 100, 150, 200 and 250nm Super Ice® Service, when necessary for shipment of heat labile products (e.g. gold conjugates, resins and some fixatives) Gold particle size will influence labelling density and selection of instrument magnification when used in immunoelectron microscopy. The tables and figures below should aid the researcher in choosing the gold particle best suited to his or her research requirements (reproduced with permission from Giberson and Demaree: The Influence of Immunogold Particle Size on Labelling Density. Microscopy Research and Technique 27:355-357, 1994). Table 1: Gold Particle Labelling Densities Gold Particle Size Mean Number Particles/µm2 Standard Deviation 5nm 10nm 15nm 20nm 30nm 746 390 227 141 167 ±105 ±84 ±40 ±26 ±18 Table 2: Percent Decrease in Labelling Density for Paired Comparisons of Gold Conjugates This Study (1) Gu & D'Andrea (2) Ghitescu & Bendayan (3#) Hansen et al. (4#) Henegouwen & Leunissen (5#) Yokota (6#) 10 to 5nm 48% . 50% 46% 56% 18% 15 to 5nm 70% 73% 81% . . . 20 to 5nm 81% . . . . 51% 30 to 5nm 78% 91% . . . 80% 15 to 10nm 42% . 62% . . . 20 to 10nm 64% . . . . 40% 30 to 10nm 57% . . . . 75% 20 to 15* 38% . . . . . 30 to 15* 26% 33% . . . . 30 to 20* none . . . . 59% 1 mean diameters 5.5, 10.0, 14.6, 20.4, 28.4 2 mean diameters not given, reported as 5, 15, 30 and 40nm 3 mean diameters 4.92, 10.1, 15.9nm 4 mean diameters not given, reported as 5 and 10nm 5 mean diameters 6.7 and 12.1 6 mean diameters not given, reported as 5, 12, 18, 28, and 38nm # data extrapolated from graphs and figures to determine percentage decreases * paired comparisons not statistically significant Table 3: Paired Comparisons of Gold Particle Size Densities Particle Size + + + + 30 + + 0 0 20 + + 0 15 + 10 + Particle Size 5 10 15 20 + = Significantly Different at 0.05% Level 0 = Not Significantly Different Labeling of the characteristic electron dense granules found in the processed cheese sample is clearly evident for 5 different sized gold conjugates. The magnification for each is identical (60,000). A: 5nm, B: 10nm, C: 15nm, D: 20nm, E: 30nm, Bar in A-E= 0.5µm Gold Conjugate Technical Information Protein Conjugate Gold Particle Size OD at 520nm Approximate Protein Concentration µg/mL Approximate Number Gold Particles/mL (15)= to the 15th power Approximate Proteins Per Gold Particle Mean Working Dilution for EM Immunoglobulin 1nm - 50 2 x 10(15) 1 1:200 Protein A 1nm - 15 2 x 10(15) 1 1:200 Streptavidin 1nm - 25 2 x 10(15) 1 1:200 - - - - - - - Immunoglobulin 5nm 3.0 36 1.7 x 10(14) 3 1:150 Protein A 5nm 3.0 12 1.7 x 10(14) 4 1:150 Streptavidin 5nm 3.0 20 1.7 x 10(14) 5 1:150 - - - - - - - Immunoglobulin 10nm 3.0 30 1.7 x 10(13) 12 1:100 Protein A 10nm 3.0 10 1.7 x 10(13) 16 1:100 Streptavidin 10nm 3.0 20 1.7 x 10(13) 20 1:100 - - - - - - - Immunoglobulin 15nm 4.0 30 5 x 10(12) 27 1:75 Protein A 15nm 4.0 10 5 x 10(12) 36 1:75 Streptavidin 15nm 4.0 20 5 x 10(12) 45 1:75 Immunoglobulin 20nm 4.0 30 2 x 10(12) 48 1:50 Protein A 20nm 4.0 10 2 x 10(12) 64 1:50 Streptavidin 20nm 4.0 20 2 x 10(12) 80 1:50 Immunoglobulin 30nm 5.0 15 8 x 10(11) 86 1:25 Notes: Gold conjugates 5nm to 30nm are packaged with 20% glycerol and can be frozen. 1nm conjugates are not packaged with glycerol and should not be frozen. EM Gold conjugates - %CV: <12% for 5nm; <8% for 10,15 & 20nm; <20% for 30nm. %Singlets: >85% for all conjugates except Protein A >95%. Guidelines for use The gold conjugates are made to the highest specification and will yield excellent results when correctly used. The guidelines discussed herein are designed to help achieve optimum results for post-embedding immunolabeling (labeling of ultrathin or semithin sections) for lelectron or light microscopy. Gold conjugates are supplied in one of the two buffers listed below. The type of buffer will determine the shelf life and storage conditions of the conjugate. Buffer A: 20mM Tris (tris-hydroxymethyl-aminomethane); 20mM sodium azide; 225mM NaCl; 1% BSA; 20% glycerol. pH 8.2 Conjugates supplied in the buffer: all EM (electron microscopy), LM (light microscopy) and BL (blotting) grade conjugates. Shelf life and storage conditions: Stable for 12 months at 4°C or for years if frozen at 25°C or below. Buffer B: 10mM Na2HPO4; 10mM sodium azide; 150mM NaCl; 1% BSA. pH 7.4 Conjugates supplied in the buffer: All 1nm (ultra small) gold conjugates Shelf life and storage conditions: Stable for 12 months at 4°C - DO NOT FREEZE The following suggestions have given good to excellent immunolabeling results at the EM level. For excellent discussion on immunolabeling strategies we recommend: Griffiths, G. (1993) Fine Structure Immunocytochemistry. Springer-Verlag, New York (Although there are a number of buffers which can be used for immunolabeling we list only two: Tris Buffer: 0.242g (20mM) Tris (tris-hydroxymethyl-aminomethane) 0.9g (225mM) NaCl ultrapure water to make 100ml. Adjust pH from 7.4 to 8.2 with 1N NaOH. Phosphate Buffer: 0.148g Na2HPO4 = 0.043g KH2PO4 = 0.72g NaCl ultrapure water to make 100ml. Adjust pH from 7.4 to 8.2 with 1N NaOH. Positive/Negative Controls: The use of a positive control is important for working out dilutions of both primary and secondary antibodies and identifying an optimum signal-to-noise ratio. The positive control will also help to establish antigenic locations and what, if any, blocking measures may be required. The use of a negative control is important in the determination of background due to the gold conjugated secondary (omit the primary antibody step and incubate with buffer only), and specificity/background due to the primary antibody (use preimmune sera or an antibody that is nonspecific for the tissue in question). Fixation: All antigens are different with respect to fixation. The use of paraformaldehyde alone (2-4%) and paraformaldehyde mixed with glutaraldehyde (0.1-0.5%) is a good place to start. It is possible, with some antigens, to follow with reduced osmium tetroxide (2% aqueous osmium mixed with and equal volume of 3% potassium ferricyanide either, buffered or unbuffered). If osmium can not be used, Berryman and Rodewald (1990, J. Histochem Cytochem 38:159-170) have an excellent protocol to achieve membrane contrast without the use of osmium. Similar results can be achieved by staining the labeled sections for 5 minutes on 2% aqueous uranyl acetate followed by 2% aqueous osmium tetroxide for the same time. We prefer to use 0.1M sodium cacodylate as the buffer at a pH between 7.2-7.4. There are many acceptable buffers and buffer combinations, however, that can be used. (See also Hyat, M.A. (1989) Principles and Techniques of Electron Microscopy: Biological Applications, 3rd Edition, CRC Press, Boca Raton, FL) Resins: The approach as to what resin to use is based on immunoreactivity. Resin-free cryosections would head the list followed by hydrophilic polar resins (i.e. Lowicryl K4M, K11M, LR White, LR Gold, Unicryl) which are partially water-soluble, hydrophobic apolar resins (i.e. Lowicryl HM20, HM23) and finally the hydrophobic epoxy resins (i.e. Epon, Araldite, Spurr's). Sample Labeling Protocol: Blocking Step: 20-30 minutes at room temperature. Primary Antibody Incubation: 30-60 minutes in duration. Can be done at room temperature or 37°C. Buffer Rinse: If done on drops, use at least 6 drops (5 minutes each drop). Gold Conjugate Incubation: 30-45 minutes in duration. Can be done at room temperature or 37°C. Rinse: In the first rinse we have found it beneficial to elevate the NaCl concentration to ~2.5M from 225mM. This appears to aid in eliminating much of the background due to ionic attraction of the negatively charged gold particle. This step is about 10 minutes followed by rinses in distilled water. Post Stain: 2% aqueous uranyl acetate for 5 minutes, rinse well in distilled water followed by 5 minutes with lead stain (25ml distilled water, dissolve one sodium hydroxide pellet then add 0.125g lead citrate).
High Performance Ceramic Adhesive
Description High Performance Ceramic Adhesive is a dispersion of Aluminium Oxide in an inorganic silicate aqueous solution. It is specially formulated for bonding and sealing ceramics, metals, and quartz for applications demanding electrical and thermal insulation at high continuous service temperature and low VOC's for ultra high vacuum. It also performs at cryogenic temperatures. It provides both low electrical and thermal conductivity. Its volume resistance is 109 ohms @RT. Its Dielectric Strength 256V per mil @RT. Surfaces to be coated should be clean but wet thoroughly to ensure good adhesion. Advantages One component system. ‐ No mixing required. Inorganic system ‐ No hydrocarbons No VOC's. High service temperature ‐ Up to 1650°C, strength improves with temperature. Low temperature capability ‐ Not effected by cryogenic temperatures. High electrical and thermal resistance. Good mechanical strength. Excellent resistance to acids and alkali. Excellent moisture resistance. Suitable for ultra high vacuum applications. Refrigeration not required. High viscosity paste ‐ viscosity can be reduced by adding water. Water soluble before cure ‐ solubility is reduced with exposure to increasing temperature. Bond integrity will depend on joint design and differential thermal expansion between substrate, sample, and adhesive paste. Typical Properties (as supplied) Filler: Alumina Binder: Inorganic Silicate Diluent: Water Colour: White Consistency: smooth, flowing paste ‐ viscosity can be reduced by adding water. Alumina content by weight: >60% Density: 2.3 g/cc Soluble in water: up to 260°C exposure Shelf life: 6 months minimum after receipt of paste ‐ can be increased by adding water and/or removing skin that can form on the top layer. Storage: Store at room temperature in tightly sealed container. Do not freeze. Application Apply adhesive paste to each surface in a thin coat using a brush, spatula or dispenser. Wet the surface thoroughly to ensure good adhesion. Maintain a uniform bond line of 2‐8 mils. Apply even pressure (clamp if possible), and wipe away excess material before drying. Good mechanical strength is achieved within a matter of minutes at room temperature. Successive coats may be applied after curing. Cure Schedule (bond time/temperature) Air set for 1 to 4 hours, then heat Cure for 2 hour cure at 93°C to achieve final electrical and mechanical properties. Blistering may occur if the glue line is too thick or heating too rapid. Strength improves with temperature and it becomes almost insoluble if exposed to temperatures above 260°C. Adhesive must be cured before use at elevated or cryogenic temperatures. Typical Properties (when cured) Recommended thickness: 2‐8ml dried (25‐100µm) Volume resistance: is 109 ohms @RT, 105 ohms @538°C Dielectric Strength: 256 volts per mil @RT, 100 volts per mil @538°C Thermal Conductivity: Torque strength: 6 ft‐lbs CTE: 7.6 x10‐6 in/in/°C Moisture Resistance: Excellent after firing above 370°C Alkali Resistance: Good after firing above 370°C Acid Resistance: Excellent after firing above 370°C
High Performance Nickel Paste
Description High Performance Nickel Paste is a dispersion of 20 µm Nickel flakes in an inorganic silicate aqueous solution. It is specially formulated for applications demanding high continuous service temperature and/or low VOC's for ultra high vacuum applications but it also performs at cryogenic temperatures. It provides good electrical and thermal conductivity. Its sheet resistance is 2.00 ohms/sq/mil (25µm). Its thermal conductivity is 2.6 W/m°K. Surfaces to be coated should be clean and free of grease. Advantages One component system. ‐ No mixing required. Inorganic system ‐ No hydrocarbons No VOC's. High service temperature. ‐ Up to 538°C, strength improves with temperature. Low temperature capability. ‐ Not effected by cryogenic temperatures but bond integrity will depend on joint design and differential thermal expansion between substrate, sample, and paste. Good electrical and thermal conductivity. Suitable for ultra high vacuum applications. Refrigeration not required. Thin paste (20,000 ‐ 25,000 cP) ‐ viscosity can be reduced by adding water. Water soluble after cure ‐ solubility is reduced the higher the temperature it is exposed to. Typical Properties (as supplied) Filler: Nickel Binder: Inorganic Silicate Diluent: Water Colour: Light Grey Viscosity: 20,000 ‐ 25,000 cP @ 25°C Consistency: smooth, flowing paste ‐ viscosity can be reduced by adding no more than 10% water by weight. Recommended thickness: 2‐8 mils (25‐100µm.) applied as a glue line or coating. Nickel content by weight: >70% Density: 2.8 g/cc Shelf life: 6 months minimum after receipt of paste ‐ can be increased by adding not more than 10% water and/or removing skin that can form on the top layer. Storage: Store at room temperature in tightly sealed container. Do not freeze. Application Mix thoroughly. Apply adhesive paste to each surface in a thin coat using a brush, spatula or dispenser. Prewet the surfaces to improve adhesion. Maintain a uniform bond line of 2‐8 mils. Apply even pressure (clamp if possible), and wipe away excess material before drying. Good mechanical strength is achieved within a matter of minutes at room temperature. Successive coats may be applied after curing. Cure Schedule (bond time/temperature) Air set for 2 to 4 hours, then heat Cure for 2 hour cure at 93°C to achieve final electrical and mechanical properties. Blistering may occur if the glue line is too thick or heating too rapid. Strength improves with temperature and it becomes almost insoluble if exposed to temperatures above 260°C. Adhesive must be cured before use at elevated or cryogenic temperatures. Typical Properties (when cured) Recommended thickness: 0.5‐1.5 mils dried (12.5‐37.5µm.) Sheet resistance: 3.5 ohms/sq/mil (25µm) after air dry. 2.00 ohms/sq/mil (25µm) after heat cure. Thermal Conductivity: 2.61 W/m°K. Soluble in water: up to 260°C exposure. Will still soften in water but may require abrasion to remove. Bond strength: Moderate but brittle.
High Performance Silver Paste
High performance silver paste is a dispersion of 20µm silver flakes in an inorganic silicate aqueous solution. It is specially formulated for applications demanding high, continuous service temperature and/or low VOCs for high vacuum applications, but it also performs at cryogenic temperatures. It provides both high electrical and thermal conductivity. Its sheet resistance is 0.08 ohms/sq/mil (25µm). Its thermal conductivity is 9.1 W/m°K. Surfaces to be coated should be clean and dry. Advantages One component system. No mixing required. Inorganic system ‐ no hydrocarbons no VOC's. High service temperature. ‐ Up to 927 °C, strength improves with temperature. Low temperature capability. Not affected by cryogenic temperatures, but bond integrity will depend on joint design and differential thermal expansion between substrate, sample and paste. Electrically and thermally conductive. Suitable for high vacuum applications. Refrigeration not required. High viscosity paste ‐ viscosity can be reduced by adding water. Water soluble after cure ‐ solubility is reduced, the higher the temperature it is exposed to. Typical properties (as supplied) Pigment: silver Binder: inorganic silicate Diluent: water Consistency: smooth, flowing paste ‐ viscosity can be reduced by adding water. Silver content by weight: >60% Density: 2.3 g/cc Shelf life: 6 months minimum after receipt of paste ‐ can be increased by adding water and/or removing skin that can form on the top layer. Storage: at room temperature in tightly sealed container. Do not freeze. Cure schedule (bond time/temperature) Achieves good mechanical strength with low conductivity in a matter of minutes at room temperature, but requires a 2 hour cure at 93°C to achieve stated high conductivity and a strong bond. Strength improves with temperature and it becomes almost insoluble if exposed to temperatures above 260°C. Must be cured before use at cryogenic temperatures. Typical properties (when dried) Recommended thickness: 0.581.5 mils dried (12.5 ‐ 37.5µm.) Sheet resistance: 0.08 ohms/sq/mil (25µm). Thermal conductivity: 9.1 W/m°K. Soluble in water: up to 260°C exposure. Will still soften in water but may require abrasion to remove. Bond strength: moderate but brittle.
High Temperature Carbon Paste
Description High Temperature Carbon Paste is a dispersion of Carbon flakes in an inorganic silicate aqueous solution. The High Temperature Carbon Paste is aimed for applications where a conductive cement is needed which withstand temperatures up to 2000°C. This paste is ideally suited for mounting specimens on hot stages for SEM, FESEM, XPS, ESCA, SIMS, AUGER systems. Also for applications where silver migration or a reaction with silver or nickel flakes could be a problem, this product would be an excellent alternative. It is specially formulated for applications demanding ultra high continuous service temperature and/or low VOC's for ultra high vacuum applications but it also performs at cryogenic temperatures. It provides moderate electrical and thermal conductivity. Its sheet resistance is 4.60 ohms/sq/mil (25µm). Its thermal conductivity is estimated to be 1 W/m°K. Surfaces to be coated should be clean and free of grease. Advantages One component system. ‐ No mixing required. Inorganic system ‐ No hydrocarbons No VOC's. High service temperature. ‐ Up to 2000°C, strength improves with temperature. Low temperature capability. ‐ Not effected by cryogenic temperatures but bond integrity will depend on joint design and differential thermal expansion between substrate, sample, and paste. Moderate electrical and thermal conductivity. Suitable for ultra high vacuum applications. Refrigeration not required. Thin paste (20,000 ‐ 25,000 cP) ‐ viscosity can be reduced by adding water. Water soluble after initial cure ‐ solubility is reduced the higher the temperature it is exposed to. Typical Properties (as supplied) Filler: Carbon Binder: Inorganic Silicate Diluent: Water Colour: Dark Grey/Black Viscosity: 20,000 ‐ 25,000 cP @ 25°C Consistency: smooth, flowing paste ‐ viscosity can be reduced by adding no more than 10% water by weight. Recommended thickness: 2‐8 mils (25‐100µm.) applied as a glue line or coating. Carbon content by weight: 50‐60% Density: 1.6 g/cc Shelf life: 6 months minimum after receipt of paste ‐ can be increased by adding not more than 10% water and/or removing skin that can form on the top layer. Storage: Store at room temperature in tightly sealed container. Do not freeze. Application Mix thoroughly. Apply adhesive paste to each surface in a thin coat using a brush, spatula or dispenser. Abrade and clean surface for best results. Preweting the surfaces may improve adhesion. Maintain a uniform bond line of 2‐8 mils. Apply even pressure (clamp if possible), and wipe away excess material before drying. Good mechanical strength is achieved within a matter of minutes at room temperature. Successive coats may be applied after curing. Cure Schedule (bond time/temperature) Air set for 2 to 4 hours, then step heat Cure for 2 hour cure at 93°C then 2 hour cure at 260°C to achieve final electrical and mechanical properties. Blistering may occur if the glue line is too thick or heating too rapid. If the second step cure is not performed then the matrix can pick up moisture. Strength improves with temperature and it becomes almost insoluble if exposed to temperatures above 260°C. Adhesive must be cured before use at elevated or cryogenic temperatures. Typical Properties (when cured) Recommended thickness: 0.5‐1.5 mils dried (12.5‐37.5µm.) Sheet resistance: 6.5 ohms/sq/mil (25µm) after air dry. 4.6 ohms/sq/mil (25µm) after step heat cure. Estimated Thermal Conductivity: 1 W/m°K. Soluble in water: up to 260°C exposure. Will still soften in water but may require abrasion to remove. Bond strength: Moderate but brittle.
Histocryl Embedding Resin
WHY HISTOCRYL It is now well established that there are many advantages in embedding in resin rather than paraffin wax. Resin causes less shrinkage and separation of tissue layers and thinner sections can be cut. Thinner sections are sharper under the microscope. Semi-thin sections are particularly useful in the diagnosis of renal disease. In the renal glomerulus, pathological changes previously requiring electron microscopy for diagnosis can now be revealed under the light microscope. The histology of densely cellular tissues benefits considerably from thin sections and this is particularly true of lymph nodes in the diagnosis and classification of lymphomas. Hard dense tissues such as bone and some botanical specimens are given improved support during sectioning, preserving the juxtaposition of hard and soft tissues. For this reason many laboratories now routinely embed bone marrow trephines in resin. Histocryl is a hydrophilic acrylic resin, simple to use and formulated specifically for light microscopists. For those laboratories currently using an acrylic resin such as HEMA glycol methacrylate or commercially branded methacrylates, no alteration need be made to their current processing schedule. Making up Histocryl We supply Histocryl uncatalysed. That material has an almost unlimited shelf-life at room-temperature and does not suffer from lengthy transportation. 100mL Histocryl plus 1.5g benzoyl peroxide = catalysed resin, however, it is recommended that the whole bottle is catalysed. Mix this well, a magnetic stirrer on low speed for about one hour is effective. Undissolved catalyst that is allowed to concentrate in the bottom of a container may generate heat and cause rapid polymerisation of the entire container. Catalysed resin keeps refrigerated for about a year. Use this catalysed medium for infiltrating the tissue and it could be used for embedding and then oven-curing. For rapid polymerisation without oven-curing, when ready to embed, add to 10mL of catalysed resin 1 drop accelerator to initiate polymerisation. The tissue will polymerise within 10-20 minutes. Smear accelerator onto the base of the mould. Add 1 drop accelerator per 10ml catalysed resin. Mix well, fill mould, add tissue. NB The moulds should be placed in a bath of ice cold water to dissipate the heat generated by the exothermic reaction. Brittle blocks may result from excessive heat produced during polymerisation. This could be due to too much accelerator and especially larger blocks, which may not have been cooled adequately during polymerisation. It is very important that blocks are cooled in ice cold water to dissipate heat produced during the exothermic reaction. The rate of polymerisation can be adjusted by varying the ratio of resin and accelerator e.g.: One drop to 10ml freshly catalysed resin (with 1.5% benzoyl peroxide paste) 10 minutes One drop to 20ml freshly catalysed resin (with 1.5% benzoyl peroxide paste) 15 minutes One drop to 25ml freshly catalysed resin (with 1.5% benzoyl peroxide paste) 20 minutes Our Peel-a-Way embedding moulds are well suited for resin embedding for histology. Fixation: Most routine fixatives can be utilised with Histocryl (neutral buffered formalin is recommended), fixation time as always, depends on the type and size of tissue. Dehydration: A graded ethanol series is the method of choice, times again are dependent on the size of the tissue. Graded acetones should not be used. A typical dehydration schedule for a block (12 x 10 x 3mm) on a mixer would be: 1. 70% alcohol - 30 minutes 2. 90% alcohol - 30 minutes Two changes absolute alcohol 30 minutes each. Infiltration: Infiltrating solution: 100ml Histocryl plus 1.5g benzoyl peroxide paste. Mix thoroughly until solution becomes clear. Infiltrate tissue in 2-3 changes of catalysed resin 60 minutes each or overnight, depending on tissue and size. When fully infiltrated the tissue becomes translucent. Cutting and Mounting: Histocryl can be sectioned using a steel knife and a standard microtome, but the method of choice is the use of a motorised microtome and glass (Ralph type) knives. Sections can be obtained from 1-5µm, floated onto a warm water bath picked up onto clean slides and dried on a hot plate at 60°C for at least 30 minutes. Most microscopist find a thickness of 1.5 or 2µm is the best compromise between sharpness and contrast. Staining: It is not necessary to etch or remove the resin before staining. Most routine stains give good results on tissue embedded in Histocryl using standard times and temperatures, although it may occasionally be necessary to extend some staining times. Mounting: For best results air dry sections prior to mounting. DPX or Canada Balsam are recommended mounting media.
JB-4 Embedding Kit
FIXATION and DEHYDRATION: Tissue can be fixed with routine light microscopy fixatives. Best results are obtained with neutralised buffered formalins or Bouins. Specimen size should be kept small at 0.2 x 0.2cm. Dehydrate samples through a graded series of ethanol 70%-95%. Because the JB-4 resin is water soluble, complete dehydration through 100% ethanol is not necessary, although recommended especially for large or dense tissue. Clearing agents such as xylene or chloroform are not necessary. Fixation, dehydration, and infiltration can be accomplished manually or automated with the use of a regular tissue processor used for paraffin processing. Processing through cold (4 degrees C) fixative, buffer rinse and infiltration resin can be used for optimal enzyme and antigen retention and preservation. This procedure uses the infiltration resin as the dehydrating agent replacing the alcohol series. No alcohol dehydration is needed, but recommended for large, bloody, or fatty tissue. PREPARATION OF JB-4 CATALYZED INFILTRATION RESIN: Prepare the infiltration resin as follows: 100ml of JB-4 Solution A add 0.90 grams of dry Catalyst C. Mix until dissolved. Careful weighing of the catalyst is necessary for correct polymerisation control. This infiltration solution may be stored for 5-6 weeks at 4°C in a dark bottle. The percentage of catalyst added to Solution A should be decreased to 0.5%-0.7% when using large quantities for automatic processor units. This aids in solution preservation and minimises heat sensitivity under processor conditions. Also, decreasing catalyst percentage (0.7%) seems useful in providing positive immunostaining. Infiltration time ranges from 2 hours to several days depending on size and tissue density. The tissue appears translucent and usually sinks to the bottom of the container. These solutions should always be kept cold. When processing for routine samples, infiltration solution should be changed 3-4 times, 30-90 minutes duration for each change. Solutions and tissue should be agitated on a rotator or hematology shaker during infiltration. EMBEDDING: Have embedding moulds, labels, ice bath, gloves, instruments and cold fresh catalysed Solution A ready before proceeding. Prepare the embedding resin as follows: Add exactly 1ml of JB-4 Solution B to 25ml of fresh catalyzed Solution A. Never use exhausted infiltration resin for embedding. Stir well and place into an ice bath while embedding to retard premature polymerisation. Anaerobic conditions are needed for polymerisation. Moulds or embedding capsules must be filled and covered or capped tightly, using EBH-2 block holders Blocks will cut easier after removing from mould by exposure to air for a few hours. Polymerisation is complete at room temperature in 50 minutes or less. NOTE: Polymerisation proceeds more rapidly in larger batches; therefore, volumes should always remain under 50ml during polymerisation. Polymerization will take somewhat longer in cold temperatures. SECTIONING: Optimal sectioning is performed with a microtome designed for plastic embedments. 0.5 micron-3 micron sections are cut with a dry glass or diamond knife, collected with forceps, and transferred onto a room temperature water bath surface, releasing sections before they touch the water. 1-2 drops of concentrated NH4OH added to the water-bath may aid in flattening sections. Sections are collected on pre-cleaned glass slides and air-dried before staining. STAINING: Dry sections are stained directly without xylene or alcohol pre-treatment. Longer staining times or higher stain concentrations may be necessary for thin sections. Alcohol or water rinses may be necessary after staining but the last step in the process should be water. Our Tissue-Tack (EMS71301-01) may be used to affix tissue to the slide during lengthy procedures. Slides are mounted while still moist with most any mounting media.
Latex Particles Suspension
CHARACTERISTICS OF POLYSTYRENE PARTICLES AJUSTING PARTICLE NUMBERS TO REQUIRED CONCENTRATIONS VOLUME, NUMBER AND SURFACE AREA VS. PARTICLE SIZE AVIDIN POLYSTYRENE PARTICLES (0.8µm) ANTIBODY COATED POLYSTYRENE PARTICLES (0.8µm) SpheroTECHNICAL NOTES SELECTED REFERENCES LATEX PARTICLES SUSPENSION CHARACTERISTICS OF POLYSTYRENE PARTICLESDensity: 1.05 Refractive Index: 1.59 Composition: Linear polystyrene Shape Uniform microspheres Porosity: Nonporous Compatibility with organic solvent: Inert to alcohol and DMSO but soluble in DMF, acetone, acetonitrile, xylene, chloroform and methylene chloride. Functional groups: Located on the surface with alkyl linker arms. Functional group contents: 0.8µm Carboxyl particles: ~ 50 ueq/g solid 0.8µm Amino particles: ~ 15-20 ueq/g solid AJUSTING PARTICLE NUMBERS TO REQUIRED CONCENTRATIONS For 1 ml of 1% w/v (10 mg) particlesDIAMETER (µm)SURFACE AREA (cm²)#(x10e9) 0.05 11428 145513 0.10 5714 18189 0.50 1143 145.51 0.80 714 35.53 1.0 571 18.19 1.5 381 5.389 2.0 286 2.274 2.5 229 1.164 3.0 190 0.6737 3.5 163 0.4242 4.0 143 0.2842 4.5 127 0.1996 5.0 114 0.1455 5.5 104 0.1093 6.0 95 0.0842 6.5 88 0.06625 7.0 81 0.0530 8.0 0.03552 10.0 0.01819 12.0 0.01052 15.0 0.00539 VOLUME, NUMBER AND SURFACE AREA VS. PARTICLE SIZEVolume of particle: V = (3.14/6) x D3 x 10e-12 cm3/particle Number of particles: N = (6W/3.14PD3) x 10e12 particles Surface area of particles: A = (6W/PD) x 10e4 cm² Total surface area of 1 ml of 5% w/v (50 mg) particle: A = 2857/D cm² Total surface area of 20 ml of 0.25% w/v (50 mg) particles: A = 2.857/D cm² Where W = Weight of polymer in gram P = Density of polymer (polystyrene = 1.05) D = Diameter of particles in micrometer AVIDIN POLYSTYRENE PARTICLES (0.8µm) Covalently coated with egg white avidinAvidin Contents: ~ 14 µg/mg solid ~ 0.212nmole/mg solid Binding capacity to Biotin-Fluorescein: ~ 0.46nmole/mg solid ANTIBODY COATED POLYSTYRENE PARTICLES (0.8µm)Antibody contents: ~ 14µg/mg solid ~ 0.2µg/sq cm ~1.5x104 IgG/particle Binding capacity to IgG-FITC ~ 4µg/mg The following Sphero TECHNICAL NOTES are available via support@proscitech.com STN-1: Particles Coating Procedures STN-2: Determination of Antibody binding to particles STN-3: Binding Capacity of Avidin Magnetic Particles STN-4: Binding Capacity of Gt-anti-Ms-IgG Magnetic Particles STN-5: Binding Capacity of Streptavidin Magnetic Particles STN-6: Binding Capacity of Biotin Magnetic Particles STN-7: Separation of Mononuclear Cells from Peripheral Blood using SPHERO Gt-anti-Ms-IgG Magnetic Particles STN-8: Calibration and Performance Tracking of Flow Cytometer Using SPHERO Calibration Particles STN-9: Measuring MESF with Flow Cytometer Using SPHERO Rainbow Calibration Particles. STN-10: Magnetic Particles Enzyme Immunoassay (MPEIA) using UltraMag Separator System (UMS-4000) Selected References Ahmed, S., El-Asser, M., Paul, G., Vanderhoff, J., "Cleaning Latexes for Surface Characterization by Serum Replacement", J. of Colloid and Interfacial Science, Vol. 73, 388 (1980) Labib, M., Robertson, A., "Application of a Diafiltration Technique in Latex Studies", J. of Colloid and Interfacial Science,, Vol. 67, 543 (1978) Hechemy, K., Michaelson, E., "Latex Particle Assay in Laboratory Medicine, Part 1", Lab Management, June/July, 27 (1984) Seaman, G., Goodwin, J., "Physiochemical Factors in Latex Rapid Agglutination Tests", Amer. Clin. Prod. Rev., June, 26 (1986) Schwenzer, K, MacCrindle, C., "Particle Concentration Fluorescence Immunoassay: Clinical Applications", Clin. Lab Prod.,Sept, 21 (1985) Jolley, M., Wang, C., Ekenberg, M., Zuelke, M., Kelso, D., "Particle Concentration Fluorescence Immunoassay (PCFIA): A New, Rapid Immunoassay Technique with High Sensitivity", J. Immunol. Methods, Vol. 67, 21 (1984) Leahy, D., Shah, D., Todd, J., "A Method for Attachment of Peptides to Solid Surface with Enhanced Immunoreactivity", BioTechniques, Vol 13 (5), 738 (1992) Leahy, D., Shah, D., Arima, T., et al., "Improved Serological Detection of Hepatitis C Virus with a Paramagnetic Microparticles Assay using Multiple Antigenic Sequences", Transfusion, Vol. 32 (6), 548 (1992) Todd, J., Kink, J., Shah, D., et al., "A novel Semi-automated Paramagnetic Microparticle Based Enzyme Immunoassay for Hepatitis C Virus: Its application to serological testing", J. of Immunoassay, Vol. 13 (3), 393 (1992) Phillips, D., Reimer, C., Wells, T., Black, C., "Quantitative Characterization of Specificity and Potency of Conjugated Antibody with Solid-Phase, Antigen Bead Standards", J. of Immunol. Methods, Vol. 34 315 (1980) Kreuter, J., Berg, U., Liehl, E., Soliva, M., Speiser, P., "Influence of the Particle Size on the Adjuvant Effect of Particulate Polymeric Adjuvants", Vaccine, Vol. 4, June, 125 (1986) Hadfield, S., Lane, A., McIllmurray, M., "A Novel Coloured Latex Test for the Detection and Identification of more than one Antigen", J. of Immunol. Methods, Vol. 97, 153 (1987) Staros, J., Wright, R., Swingle, D., "Enhancement by N-Hydroxysulfo-Succinimide of Water-Soluble Carbodiimide Mediated Coupling Reagents", Anal. Chem. Vol. 156, 220 (1986) Van den Hul, H. Vanderhoff, J., "The Characterization of Latex Particle Surface by Ion Exchange and Conductiometric Titration", J Electroanal. Chem. Vol. 37, 161 (1972) Andrew J. Beavis and Kenneth J. Pennline, "Detection of Cell-Surface Antigens Using Antibody-Conjugated Fluorospheres (ACF): Application for Six-Colour Immunofluorescence", BioTechniques 21:498-503 (September 1996)
Liquid Blocker Pen
Usage of the Liquid Blocker Pen: The Liquid Blocker Pen is most useful for immunity staining (PAP method, ABC method, etc.) and fluorescent antibodies method in carrying out the maximum efficiency with minimum amount of extremely valuable antiserum. Method for Paraffin Sections: Sections should be quickly deparaffinised by xylene, then washed out with ethanol; dry or wipe away excess liquid around the section on the glass slide with tissue paper. Encircle the section on a glass slide with the Liquid Blocker Pen as shown below in the illustration, and dry the circle for 1-2 minutes at room temperature; then soak the section in a phosphoric acid buffer solution (PBS or TBS), for about 5 minutes. Note: The Liquid Blocker Pen makes a water repellent circle which avoids the need to wipe around the section after every staining. Circle made by the Liquid Blocker Pen is removable by xylene. Since it is not influenced by water-soluble mounting media, there is no interruption in microscopic observation. This water repellent Liquid Blocker Pen is applicable not only to immunity staining but also to ASD staining and/or Enzyme staining. Method for Frozen Sections: Glass slides must be cleaned by xylene and ethanol or coated with egg-albumin glycerine or poly-L-lysine before the Liquid Blocker Pen is applied. Note: If the surface of the glass slide is treated with hydrophile, it is necessary for the glass slide to soak in a 0.1N hydrochloric acid solution for about 30 seconds and then dry it after washing it with water in order to prevent the circle from disappearing.
Lowicryl Resin
Lowicryl K4M/K11M and Lowicryl HM20/HM23 user note covering fixation, dehydration at low temperatures, preparation of resins & infiltration, polymerisation, sectioning, staining of sections, and Cytochemistry and Immunocytochemical labelling. Lowicryl K4M/K11M and Lowicryl HM20/HM23 1. INTRODUCTION: Lowicryl K4M and HM20 are highly cross linked acrylate-and methacrylate-based embedding media which have been designed for use over a wide range of embedding conditions (3,8,14). These resins have been formulated to provide low viscosity at low temperatures: K4M is usable to -35°C, and HM20 to -70°C. The investigator also has a choice of either a polar (hydrophilic; K4M) or a non polar (hydrophobic; HM20) embedding medium (8). Both resins are photopolymerised by long wavelength (360nm) ultraviolet light. Since the initiation of the polymerisation is largely independent of temperature, blocks may be polymerised at the same temperatures which are used for infiltration. The resins may also be chemically polymerised at 60°C. The hydrophilic properties of K4M provide two distinct advantages. During dehydration and infiltration the specimens may be kept in partially hydrated state, since K4M may be polymerised with up to 5% (by weight) water in the block (8,14). Secondly, K4M is particularly useful for immunolabeling of sections using specific antisera or lectins (see section 8). The use of K4M results in a better structural preservation (21), an improved preservation of antigenicity (7,20,22) and a significantly lower background labelling. K4M and HM20 have also been used to produce high contrast images of completely unstained thin sections in the scanning transmission electron microscope by Z-contrast(10). K4M and HM20 are usable at room temperature as well as low temperatures; the applications of the resins are left to the discretion of the investigator. This booklet mainly addressed the techniques of low temperature embedding, solutions to the most common problems which are encountered in low temperature work, and general suggestions for the use of K4M and HM20. Lowicryl K11M and HM23 have similar properties K4M and HM20 but can be applied at 20°C to 30°C lower temperature. K11M has in addition a much lower viscosity compared with K4M. The two resins have been designed to explore freeze substitution combined with low temperature embedding (below -50°C). They can obviously also be used in the same way as K4M and K11M (2,9). 2. FIXATION: Any of the standard aldehyde fixation procedures (perfusion, immersion or combinations there of) may be used. Since the resins are in most cases photopolymerised, the use of fixatives which also have staining properties (e.g. osmium tetroxide) is not generally recommended. An excessive staining of the material will interfere with the penetration of UV light into the centre of the specimen, resulting in a incomplete polymerisation (1,3,8). Excess osmium tetroxide in the specimen will also attack the unsaturated bonds in the resin. Naturally occurring pigments, if present in usual amounts, generally do not interfere with the polymerisation of the Lowicryl resins. Some samples which are heavily pigmented and absorb strongly at 360 nm may produce blocks of less than optimal quality (1). To insure the adequate penetration of UV light and an even polymerisation, individual samples should be <0.5mm3. Tissues, cell pellets, etc. May be minced either in the later stages of fixation or in the buffer wash immediately following fixation. 3. DEHYDRATION AT LOW TEMPERATURES: The majority of low temperature embedding procedures are performed in one of two ways: (1) gradually decreasing the temperature during dehydration, as the material is exposed to an ascending series of concentrations of the dehydrating agent, or (2) freeze-substitution. For most routine applications, the first method is recommended. 3.1 The Progressive Lowering of Temperature (PLT) Technique: This procedure involves step-wise reductions in temperature as the concentration of dehydration agent is increased (3,8,14). A temperature is selected at each step which is above the freezing point of the concentration used in the step just before; this is indeed the concentration of the dehydration agent contained in the tissue block, when introduced into the next higher concentration of the dehydrated series. For the freezing points of various dehydrating agents, consult graph 1. Graph 1: Freezing points of commonly used dehydrating agents (solvents) as a function of concentration. EGOH Ethylene gylcol. Note the rise in the freezing point of EGOH at higher concentrations. DURING DEHYDRATION AND FILTRATION, THE SAMPLES SHOULD BE PERIODICALLY AGITATE EITHER BY STIRRING WITH A TOOTHPICK OR BY GENTLY SWIRLING THE SAMPLE VIALS. Most polar and non-polar dehydrating agents may be used with both resins. Due to its hydrophobic nature, however, HM20 is immiscible with ethylene glycol and dimethylformamide. Both resins are freely miscible with methanol and ethanol. A representative dehydration schedule for ethanol is given as follows: Ethanol Temperature Time Vol. % K4M HM20 K11M HM23 min. 30 0°C 0°C 0°C 0°C 30 50 -20°C -20°C -20°C -20°C 60 70 -35°C -50°C -50°C -50°C 60 95 -35°C -50°C -60°C -60°C 60 100 -35°C -50°C -60°C -80°C 60 100 -35°C -50°C -60°C -80°C 60 The times and temperatures above the minimum values and have to be adjusted accordingly to the type of specimen and its size. Schedules for other solvents can be developed provided that their freezing points are considered. (See graph 1). 3.2 Achieving Low Temperature: There are several methods to achieve the low temperatures for dehydration, infiltration and polymerisation (11): Balzers Low Temperature Embedding (LTE) Apparatus. Provides four sample holding blocks, which may be preset to any temperature from 0°C to -50°C. Also contains a stirring head for continuous sample agitation. Information on this apparatus is available from Balzers Corp. For -20°C, use ice: NaCl, 3:1 (wt:wt). Monitor the temperature carefully, as this mixture requires periodic replenishment. The lifetime of the mixture may be lengthened by keeping it in a glass lined Dewar, and by placing the Dewar in a refrigerator or a cold box. To minimize temperature gradients, it is preferable to use an aluminium block with drilled holes which will accommodate the sample vials. The metal block is first placed in the cooling bath and allowed to equilibrate before the sample vials are placed in the block. For temperatures of -35°C to -40°C, a household chest-type freezer may be used. For lower temperatures (i.e., -30°C to -70°C) use either a low temperature chest-type freezer or mixtures of o- and m-xylene in combination with crushed dry ice¹. For xylene mixtures, refer to Graph 2. The temperature is determined by the volume ratios of o- to m-xylene. Crushed dry ice is added to the xylene mixtures to form a thick slurry. When mixed in a Dewar flask, these xylene-dry ice mixtures will maintain a constant temperature for ca. 8-10 hours. Graph 2: Temperatures of crushed dry ice-xylene slurries, as a function of the ration of o- to m-xylene. XYLENE VAPORS ARE TOXIC. WORK WITH XYLENE COOLING BATHS ONLY IN A WELL-VENTILATED FUME HOOD. 3.3 Freeze-Substitution: The Lowicryl resins have been successfully used with freeze-substitution methods. The primary advantage of these techniques is that the infiltration and polymerisation temperatures need to be raised above -35°C. These techniques require special apparatus, since a high initial rate of cooling is critical for such procedures. For details on instrumentation contact the following firms or their distributors: Propane Jet Freezer: Balzers Corporation Cryoblock Liquid Helium Freezer: Reichert-Jung. For details on the techniques involved, consult references (12) and (17). 4. PREPARATION OF RESINS AND INFILTRATION: METHACRYLATES, SIMILAR TO OTHER EMBEDDING MEDIA, MAY CAUSE ECZEMA ON SENSITIVE INDIVIDUALS. ALWAYS USE GLOVES FOR ALL STEPS INVOLVING USE OF THE RESINS. IN CASE OF CONTACT WITH SKIN, WASH THOROUGHLY WITH SOAP AND WATER. SUITABLE GLOVES WITH GOOD PROOFNESS TO ACRYLATES ARE THE "KIMGUARD VINYL GLOVES" OF KIMBERLY-CLARK. 4.1 Mixing Instructions: Due to their very low viscosities, the Lowicryls do not require vigorous stirring to mix the resin components. Mixing too vigorously or for prolonged periods may result in the incorporation of oxygen into the resin, thereby interfering with the polymerisation. This is especially important if a thermal polymerisation ( 60°C) is used. Avoid inhaling the vapours from the resins. Use a well-ventilated fume hood for mixing the Lowicryls. Weigh out, into a tared vial, the cross-linker and the monomer. Mix gently by one of the following methods for three to five minutes: Bubble a continuous stream of dry nitrogen gas into the mixture with a Pasteur pipette. The nitrogen stream will mix the resin, and at the same time it will prevent the incorporation of oxygen. Mix gently with a glass rod. If the vial has a snap-cap or lid, slowly rock the covered vial from side to side, avoiding the formation of air bubbles or foaming. Add the initiator, and continue mixing until the initiator is completely dissolved in the resin. 4.2 Mixtures for Ultraviolet Polymerisation: K4M. HM20 HM20 Cross-linker A 2.70gm Cross-linker D 2.98gm Monomer B 17.30gm Monomer E 17.02gm Initiator C* 0. 10gm Initiator C* 0.10 gm K11M HM23 Cross-linker HM20 1.0g Cross-linker F 1.1g Monomer I 19.0g Monomer G above -50 °C 18.9g Initiator C 0.1g Initiator C -50°C to -70°C 0.1g Initiator J below -70°C 0.1g Initiator J 0.15g *For polymerisation from -50°C to 0°C. Above 0°C, the initiator C should be replaced by the same amount of benzoin ethylether. The above mixtures will produce blocks of average hardness. The hardness may be varied by incorporating more or less cross-linker to resin mixture (more cross-linker produce harder blocks). For HM20 , the cross-linker concentration may be varied from 5 to 17 weight % (1.0 to 3.4 gm/20gm resin). For K4M, the cross-linker concentration may be varied from 4 to 18 weight % (.08 to 3.6gm/20gm resin). 4.3 Mixture for Thermal (Chemical) Polymerisation of HM20 and K4M at 60 degrees C Although the resins are primarily designed for UV polymerisation, it is also possible to polymerise them with a more classical thermal ( 60 degrees C) technique. For such a procedure, mix a cross-linker and monomer as previously mentioned (Section 4.1). nitrogen bubbling is the method of choice. Instead of initiator C, substitute the following amounts of dibenzoyl peroxide: HM20: 0.5% (by weight) dibenzoyl peroxide K4M : 0.3% (by weight) dibenzoyl peroxide Dibenzoyl peroxide is generally supplied as a paste with dibutylphthalate or a powder moistened with water. Compensate for the added ingredients, so that the resin receive the above amounts of peroxide, exclusive of the additives. 4.4 Infiltration at Low Temperatures: Infiltration with Lowicryl resins at low temperatures is similar to room temperature infiltration with other embedding media. The exact protocol will depend upon the temperatures and dehydrating agent chosen, and the viscosity's of the dehydrating agent and of the resin at those temperatures. Typical infiltration schedule with ethanol, is given below. RESIN:ETHANOL TIME MINIMAL TEMPERATURES vol. : vol. time K4M HM20 K11M HM23 1:1 60 min. Very -50°C -60°C -80°C 2:1 60 min. viscous -50°C -60°C -80°C pure resin 60 min. below -50°C -60°C -80°C pure resin overnight or 4-16 hrs. -40°C -50°C -60°C -80 °C It is important to keep the samples in movement during infiltration in order to facilitate equilibration of the tissue interior with the bulk of the infiltration liquid. 5. POLYMERISATION: 5.1 Ultraviolet Polymerisation at low temperature The Polymerisation Chamber: Samples may be polymerised in either BEEM or gelatin capsules. A suitable capsule holder is required so that the capsules receive UV irradiation from all sides. A stand (fig.1) is constructed from heavy gauge wire, and finer gauge twisted wire loops are soldered onto the stand to hold the capsules. The size of the capsules is important, large volumes, over 1ml, can easily lead to a temperature increase during polymerisation. The heat produced by the exothermic polymerisation reaction is not dispatched to the surrounding. The same happens when the samples are polymerised too fast (for further details see ref.4). Fig. 1. A wire capsule holder for UV polymerisation. The light source must be 360 nm long-wave length UV, preferably two 15-watt fluorescent tubes, similar to those used for thin layer chromotography ². ² Philips TLD 15W or similar fluorescent tube. A polymerisation chamber (Fig. 2) can be constructed which will fit in a deep chest-type freezer or in a cold room. To provide diffuse illumination, a right-angle reflector is suspended below the UV lamps. All six inner surfaces as well as the reflector, should be lined with aluminium foil. The capsule holder is placed 30 - 40 cm below the fluorescent lamps. The entire box should be too tightly constructed; ventilation from the top and bottom will provide air circulation and will minimize temperature gradients in the chamber. Fig 2. A polymerisation chamber for indirect UV irradiation, the UV source (1) is diffused by a right-angle reflector (2). The capsule holder (3) is place 30-40 cm below the UV source. See section 5.1 A small, hand held UV lamp may also be used, provided it emits at 360 nm. Some of the small "mineral lamps" have both a long - and short-wave UV source. In such a case, mask the dimensions of the polymerisation chamber, and reduce the lamp-to-capsule distance to ca. 10-15 cm. Irradiation from the bottom of the chamber will reduce the attenuation of the UV light by the resin. This is recommended for low-intensity UV sources. In either the large or the small polymerisation chamber, make a trial run by polymerising pure resin in capsules. Shrinkage and deformation along the sides of the block indicate that the polymerisation is too rapid. In such a case increase the distance between the lamps and the capsules. 5.2 Ultraviolet Polymerisation at Low Temperatures: Protocol: Fill capsules with fresh pre-cooled resin. The capsules should be fill to the top, to minimise dead air space over the resin. Transfer samples to the capsules with Pasteur pipettes; close the capsules, and allow them to equilibrate at the chosen temperature for 10-15 minutes. To minimise the condensation of water and the crystallisation of ice on the sample vials and capsules, all apparatus should be pre-cooled, and steps 1 and 2 should be performed in the cold. Polymerise K4M and HM20 for at least 24 hours under UV-light, at -30°C to -40°C (the lowest recommended temperature for polymerisation is -50°C. Polymerise K11M and HM23 for at least 5 days at not lower temperature than -60°C and -80°C respectively. At these low temperatures it can be difficult to keep the UV- lamps burning with sufficient yield of irradiation. This leads to a much longer polymerisation time. Remove the capsules from the cold, and continue "curing" under UV for 2-3 days at room temperature. 5.3 Chemical Polymerisation of K4M and HM20 at 60°C: Place fresh resin (see Section 4.3) in gelatin capsules; transfer samples to capsules, and fill capsules approximately 3/4 full. Close capsules and polymerise at 60 degrees C for 2-3 days. The use of gelatin capsules is recommended for chemical polymerisation at 60 degrees C. The plasticiser in BEEM capsule may interfere with polymerisation at the periphery of the blocks. The chemical polymerisation of K4M and HM20 with peroxides is an exothermic reaction. To prevent an uncontrolled rise in temperature. The capsules should be in contact with a heat sink. Use an aluminium block with predrilled holes which will accommodate the capsules (6). The capsules should fit firmly in the holes. 6. SECTIONING: For best results, trim the final pyramids with glass knives on the microtome or on a trimming apparatus. The sides and the face should be clean, and under illumination they should be clear and transparent. Trim the sides of the pyramid at an angle of 28-30 degrees from the face. The Lowicryl resins are highly crosslinked methacrylates. When they are of the correct hardness they are easily sectioned with either glass or diamond knives. K4M and K11M are hydrophilic resins. Therefore, as with other polar (water-miscible) resins, precautions should be taken to insure that the block face does not become wet during sectioning. This is best accomplished by sectioning with a level of fluid in the trough which is slightly below normal. In such a situation, the reflection from the trough fluid along the knife edge will be slightly darker than the normal bright silver colour. However, do not lower the trough fluid so much that the knife edge becomes dry. This is particularly important with diamond knives, due to the hydrophobic nature of most diamond knife edges. The most suitable procedure with diamond knives is to orient the trimmed block with the knife edge before the trough is filled. The specimen arm of the microtome is places in its lowermost position, and the trough is overfilled to form a "reverse meniscus" along the knife edge. Leave the knife in this position for 10-15 minutes. Immediately before sectioning, lower the level of the trough fluid to produce a dark silver reflection along the knife edge. Make the final advance of the knife and /or block, and commence sectioning. Since K4M and K11M are hydrophilic resins, the sections should be collected as soon as possible after they are cut. Sectioning speeds of 2-5mm/sec are recommended. Further details on sectioning are given in Lowicryl Letters No. 2. 7. STAINING OF SECTIONS: HM20 sections of completely unstained (aldehyde fixed) material give sharp images in the scanning transmission electron microscope (STEM) in the Z-contrast mode. In conventional transmission microscopy, however, the surface relief on the sections contributes to a low contrast and a lack of resolution (7). Therefore, for conventional imaging the sections must be stained. Due to the hydrophilic and hydrophobic properties of the resins, there are significant differences in the staining behavior of the resins. Also, the amount of staining and contrast which is required is to a great extend dependent upon the investigator, the techniques of staining, and the applications for which the resins are used. Therefore, only general guidelines are given here. Sections may be stained with either saturated aqueous or alcoholic solutions of uranyl acetate. Both Reynolds' lead citrate (18) and Millonig's lead acetate (15) give good results. A series of experiments (w. Villinger, unpublished data) has shown that a particularly useful combination is a first staining with saturated aqueous uranyl acetate, followed by lead acetate according to Millonig"s (15) second method: Staining at Room Temperature : HM20 HM23 K4M K11M 1. Uranyl acetate, saturated aqueous solution 35 min. 5-10 min. 2. Millonig's lead acetate . 1-3 min 1-3 min. As with all staining procedures wash well between the uranyl and lead stains, and take precautions, against carbon dioxide during the lead staining and the rinsing after the stain. This is particularly important for lead acetate staining. ³ Since K4M and K11M are hydrophilic, the sections should be incubated on drops of the stains for short periods of time. Prolonged staining may cause distortions and contamination of the sections. 8. CYTOCHEMICAL AND IMMUNOCYTOCHEMICAL LABELLING WITH K4M : K4M has been used with success in cytochemical and immunocytochemical studies, most notably in conjunction with colloidal gold particles as an electron-opaque marker. Significant improvements in structural preservation and in lower background labelling (21) are found with K4M. Colloidal gold particles may be coated with protein A (20). Sections of K4M - embedded material are first incubated with a specific antibody, and this is followed by an incubation with the protein A-gold complex. The gold particles localise in the antibodies from the first incubation, since protein A binds specifically to the region of IgG. A technique has also been described in which colloidal gold is coated with enzymes, and the substrate is localised by an incubation of thin sections on the enzyme-gold complex. Colloidal gold has also been directly coated with antibodies, tetanus and cholera toxins, and lectins. For details of these procedures, consult references (5-7, 11, 13, 19-22). 9. REFERENCES: Acetarin, J.-d. and Carlemalm, E. (1982) The chemical polymerisation of Lowicryls. In: Lowicryl Letters No. 1 Chemische Werke Lowi GmbH, Postfach, D-8264 Waldraiburg, Federal Republic of Germany. Acetrarin, J.-D., Carlemalm, E. And Villinger, W. (1986) Developments of new Lowicryl resins for embedding biological specimens at even lowr temperatures. J. Microsc. (In press) Armbruster, B.L., Carlemalm, E., Chiovetti, R., Garvito, R.M., Hobot, J.A., Kellenberger, E. And Villinger, W. (1982) Specimen preparation for electron microsocpy using low temperature embedding resins. J. Microsc. 126, 77-85. Ashford, A. Et al. (1986) in press Bendayan, M. (1981) Ultrastructural localization of nicleic acids by the use of enzyme-gold complexes. J. Histochem. Cytochem. 29, 531-541. Bendayan, M. And Orstravic, T.B. (1982) Immunochemical localization of kollikrein in the rat exocrine pancreas. J. Histochem. Cytochem. 30, 58-66. Bendayan, M. And Shore, G.G. (1982) Immunocytochemical localization of mitochondrial proteins in rat hepatocyte. J. Histochem. Cytochem. 30, 139-147. Carlemalm, E., Garvito, R.M. and Villinger, W. (1982) Resin development for electron microscopy and an analysis of embedding at low temperature. J. Microsc. 126 123-143 Carlemalm, E. Villinger, W., Hobot, J.A., Acetarin, J.D. and Kellenberger, E. (1985) Low temperature embedding with Lowicryl resins: two new formulations and some applications. J Microscopy140, 55-63. Carlemalm, E. And Kellenberger, E. (1982) The reproducible observation of unstained embedded cellular material in thin sections: visualization of an integral membrane protein by a new mode of imaging for STEM, EMBO J. 1, 63-67 De Mey, J, Moermans, M., Guens, G., Nuydens, R. And DeBrabander, M. (1981) High resolution light and electron microscopic localization of tubulin with IGS (immuno gold staining) method. Cellular Bio International Reports 5, 889-899. Escaig, J. (1982) New instruments which facilitate rapid freezing at 83K and 6K J. Microsc 126, 221-229 Horisberger, M (1979) Evaluation of collodial gold as a cytochemical marker for transmission electron microsocopy. Bio. Cellulaire 36, 253-258. Kellenberger, E. Carlamalm, E., villinger, W., Roth, J. And Garavito, R.M. (1980). Low denaturation embedding for electron microscopy of thin sections. Chemische Werke Lowi GmbH, Postfach, D-8264 Waldkraiburg, Federal Republic of Germany. Millonig, G. (1961). A modified procedure for lead staining of thin sections, J. Biophysic and Biochem. Cytol. 11, 736-739 Montesano, R., Roth, J., Robert, A. and Orci, L. (1982) Noncoated membrane invaginations are involved in binding and internalization of cholera and tetanus toxins. Nature 296, 651-653 Muller, M. Marti, T. and Kriz, S. (1980) Improved structural preservation by freeze-substitution. In: Proc. 7th European Congress on Electron Microscopy 2, 720-721. Reynolds, E.S. (1963) The use of lead citrate at high pH as an electron opaque stain in electron microscopy. J. Cell biol. 17, 208-213. Roth, J. (1982a) New approaches for in situ localization of antigens and glycoconjugates on thin sections: the protein A-gold (PAG) technique and the lectin-colloidal gold marker system. 10th International Congress of Electron Microsocpy, Hamburg (abstract). Roth, J. (1982b) The protein A-gold (PAG) technique. Qualitative and quantitative approach for antigen localization on thin sections. In: Techniques in Immunocytochemistry, Vol. I Academic Press, London pp104-137. Roth, J., Bendayan, M., Carlemalm, E., Villinger, W. And Garavito, R.M. (1981) Enhancement of structural preservation and immunocytochemical staining in low temperature embedding pancreatic tissue. J. Histochem. Cytochem. 29, 663-671. Roth, j., Berger, E.G. (1982) Immunocytochemical localization of galatosyltransferease in HeLa cells: codistribution with thiamine pyrophoshatease in trans golgi cisternae. J. Cell Biol. 93, 223-229 References to Immunolabelling: Roth, J. The Colloidal Gold Marker System for Light and Electron Microsocpy. Theory and Application. In: "Techniques in Immunocytochemistry" (eds. E.R. Bullock u. P. Petrusz) Academic Press, London vol. II, 1983 Roth, J. Brown, D. And Orci, L. Regional distribution of N-acetyl-D-galactosamine residues in the glomerular podocytes. J. Cell Biol. 96, 1189-1196, 1983 Roth, J. Application of lectin-gold complexes for electron-microsocpic localization of glycoonjugates on thin sections. J. Histochem. Cytochem. 31, 987-999, 1983 Roth, J. Application of immunocolloids in light microscopy. Preparation of protein A-silver and protein A-gold complexes and their application for localization of single and multiple antigens in paraffin sections. J. Histochem. Cytochem., 30, 691-696, 1982 Roth, J. Applications of immunocolloids in light microscopy II. Demonstration of lectin-binding sites in paraffin sections by the use of lectin-gold or glycoprotein-gold complexes. J. Histochem. Cytochem., 31, 547-552, 1983 Norman, A.W. Roth, J., and Orci, L., The vitamin D endocrine system: Steroid metabolism, hormone receptors and biological response (calcium binding proteins). Endocrine Red. 3, 331-366, 1983 Roth, J., Light and ecletron microscopic localization of antigenic sites in tissue sections by the protein A-gold technique. Acta histochem. Suppl. in press 1983 Roth, J., The preparation of protein A-gold complexes with 3nm and 15 nm gold particles on their use in labelling multiple antigens on ultrathin sections. Histochem. J., 791-801, 1982 Roth, J., Brown, D., Norman, A.W. and Orci, L Localization of vitamin D dependent calsium binding protein in mammalian kidney. Am. J. Physiology., F243 to F252, 1982 Roth, J., Thorens, B. Brown, D., Baetens, D, Garcia-Serguira, L.M.,Norman, A.W. and Orci, L., Immunochemical localization of vitamin D-dependent calcium binding protein (CaBP) in duodenum, kidney, brain and pancreas. In: Vitamin D, Chemical, Biochemical and Clinical Endocrynology of Calcium Metabolism" (eds. A.W. Norman, K4M. Schaefer, D. V. Herrath and H.-G. Grigoleit) Walter deGruyter & Co., Berin, New York pp 209-214, 1982 Thorens, B., Roth, J., Norman A.W. Perrelet, A., and Orci, L. Immunocytochemical localization of the vitamin D-dependent calcium binding protein in check duodenum. J. Cell Biol. 94, 115-122, 1982 Roth, J., and Binder M., Colloidal gold, ferritin and peroxidase as markers on electron microscopic doubling labelling lectin techniques. J. of Histochem and cytochem. 26, 163-169. (1978) Roth, J., Bendayan, M., and Orci, L., Ultrastructural localization of intracellular antigens by the use of protein A-gold complex. J. Histochem. Cytochem., 26, 1974-1981 (1978) Paiement, J., and Bendayan, M. Localization of RNA in incubated rat liver nuclei. J. Ultrastruc. Res., 81, 145-157 (1982)
LR Gold Processing & Polymerisation
Processing and polymerisation TISSUE Tissue samples up to 5 x 5 x 5mm have been successfully processed using the following schedule. However, it is recommended that tissue specimens of maximum 3 x 3 x 3mm are used and as a general rule the smaller the specimen the more efficient the impregnation. The thickness of the tissue is particularly important when polymerising darkly coloured tissue such as liver and spleen, because complete polymerisation depends on the blue light from the light source being able to penetrate the full thickness of the tissue. Tissue used is fresh and unfixed. PROCESSING Processing is performed on a rotary agitator in 10mL vials with tight fitting lids. The fluids involved are maintained in bulk at the sub-zero temperatures required. Also, it must be remembered that the final resins are sensitive to prolonged light exposure and are therefore stored in the dark and handled as infrequently as possible. We recommend the use of polyvinyl pyrrolidine to protect unfixed tissue from osmotic changes during processing. We have used PVP with an approximate molecular weight of 44,000. This can be dissolved in methanol, water and the London Resin Gold monomer. Concentrations of 50% w/v are possible in the methanol mixtures, however, at low temperatures the resulting viscosity is impractical. The following schedule shows the PVP concentrations recommended, the resulting LM work being very satisfactory. It must be said, however, that the addition of PVP in different concentrations may further improve morphology especially in the EM. FRESH TISSUE 50% methanol 20% PVP 0°C 15 min 70% methanol 20% PVP -25°C 45 min 90% methanol 20% PVP -25°C 45 min 50% LR Gold monomer/50% methanol 10% PVP -25°?C 30 min 70% LR Gold monomer/30% methanol 10% PVP -25°?C 60 min 100% LR Gold monomer -25°C 60 min 100% LR Gold monomer initiator -25°?C 60 min 100% LR Gold monomer initiator -25°?C overnight 100% LR Gold monomer initiator -25°?C 20-25 hour POLYMERISATION The addition of a light sensitive initiator is needed in order to polymerise the resin and we recommend BENZIL, an alpha-diketone, at a concentration of 0.1% w/v. The principle is shown in the diagram. The gelatin capsules are 00 size and the plastic support is a modified heamagglutination tray. The bulb involved here is a Thorn projector lamp (A?/209 FDX, 12V 100W). We have found that 7 to 9V will cause solidification within 24 hours. Like many acrylic resins oxygen will inhibit polymerisation, therefore the capsules are filled completely and lids fitted. Paper labels can be inserted into the capsules. 9 capsules (3 x 3) may be polymerised at any one time. If the upper surface is still soft after 24 hours this can be trimmed off or hardened in daylight for a few hours prior to peeling off the gelatin. The blocks once polymerised need not be stored cold, however it may prolong the activity of some enzymes to do so. The enzyme histochemistry performed to date using the resin has involved conventional reagents, times of reaction and temperatures (see Thompson and Germain, Histochemical Journal Vol.15, No. 12, December 1983.) For room temperature polymerisation, using a peroxide/amine cure, add to pure LR Gold either 1% of dry benzoyl peroxide or, more safely, 1.5% of benzoyl peroxide paste (60% in dibutyl phthalate). Infiltrate with pure LR Gold solution adding only the peroxide mix prior to polymerisation. To reduce curing exotherm, cool mould in ice water. To accelerate cure, add 1 drop of LR White accelerator to 20mL LR Gold resin/benzoyl peroxide mixture. To U.V. light cure LR Gold, add benzoin methyl ether. The precise concentration will depend on the power and emission spectrum of your UV lamp. However, a useful starting concentration would be 0.5%. The cross link density of the final resin is important. If stains are not penetrating sufficiently quickly, reduce the benzil concentration rather than the light intensity or exposure time. SECTIONING AND MOUNTING Following polymerisation, the LR Gold blocks can be stored, handled and cut at room temperature. Cutting should preferably be done using a motorised microtome and glass knife. Sections may be cut dry, picked up and placed free-floating into incubating medium or buffer wash for enzyme histochemistry or immuno-cytochemistry. It is not advisable to mount sections onto slides before reacting, since this involves heat and would be deleterious to the unfixed proteins. The section can of course be mounted in the usual way after enzyme histochemistry or immunocytochemistry has been carried out.
LR White Embedding Kits
C025 & C026 LR White Resin (1981 is an aromatic acrylic resin mixture which was introduced following the work by Causton, Gillett and Germain, 1980). It is a very low viscosity (8 cps), low toxicity, beam stable resin. It is a hydrophilic embedding medium and the sections of polymerised LR White resin are hydrophilic. This character allows immuno cytochemistry reagents to easily penetrate into the section without the need of etching, (etching sections can effect delicate tissue antigens). LR White can be polymerised by four different methods: Heat: 60 – 65°C UV irradiation (365 nm wavelength) Chemical with accelerator Microwave EMS supplies LR White in either the premixed form - ready to use (Catalysed LR White), which makes it convenient for the end user or the non-catalysed version with 9.9g of Catalyst which accompanies the LR White resin with instructions showing you how to catalyse the resin prior to use. (Usually it takes 12 hours at room temperature and 12 hours in the refrigerator for catalysed LR White resin to take effect prior to use). LR White (catalysed) should be kept at 4°C or lower. In our experience, the shelf-life of LR White is a minimum of one year @ 4°C, and up to 15 months if stored as low as -80°C. If the resin is frozen, allow it to thaw and reach room temperature prior to use. If the viscosity of the LR White resin changes (becomes thicker), stop and do not use it. LR WHITE FOR ELECTRON MICROSCOPY PROTOCOLS Fixation Conventional EM/LM: 2 to 4 hours - recommended Paraformaldehyde (3 – 4%) solution in 0.1M sodium phosphate buffer, pH 7.2. Avoid using Glutaraldehyde alone or Karnovsky's glutaraldehyde/formaldehyde mixture, for this may lead to a patchy stain or some stains not working well. For Microwave: Place specimen cubes (~1 to 1.5 mm2) in 1.5ml centrifuge tube with ~600 µl 4% Paraformadehyde in Sodium Phosphate buffer pH 7.2. Microwave irradiate the vial for 40 seconds in a cold spot. Let the vial + tissue + buffer sit for an additional 5 minutes allowing for the cooling of the buffer ≤20°C, and then irradiate again the vial + tissue + buffer for another 40 seconds. For Immunocytochemistry: 2-3 hours – Recommended 4% paraformaldehyde/0.05% Glutaraldeyde EM Grade/0.2% picric acid in 0.1M sodium Phosphate buffer, pH 7.3 [Smogyi, P and Takagi. H (1982). Note: The use of picric acid-paraformaldehdye-glutaraldehyde fixative for correlated light and electron microscopic immunocytochemistry. Neurosciences 7, 1779 – 1783.] Buffer rinses are needed at least 2 times and 30 minutes each, for all of applications. Except for the microwave, 2 rinses 3 minutes each is done outside the microwave. Post Fixation Conventional: 1 – 2% osmium tetroxide in water or buffer for one hour For dual LM/LM: post fixation with osmium tetroxide should be avoided. For Immunocytochemistry: post-osmication or “block staining” should not be carried out. However, post fixing the tissue with 1% tannic acid is recommended. Then the “Addition of phosphotungstic acid (PTA) to ethanol for dehydration step, will improve both the ultrastructure and antigenicity of pituitary tissue embedded in LR White.” [Yoko Sakai, Masahiro Hosaka, et al. Histology and Cytology, Vol.68 (2005), No. 5 p.337-34] Or, post fix in the mix equal volumes of 3% potassium ferricyanide and 2% osmium tetroxide, with a final concentration of 1.5:1 - buffered or un-buffered, to improved the contrast without using osminum tetroxide [Berryman & Roddewald, 1990. Histochem/cytochem. 38:159 – 70; Berryman et al., 1992. J. Histochem/Cytochem. 40:845-57] For Microwave: same procedure as the fixation for microwave above. Dehydration / Resin Infiltration For best results – the use of a rotator for this step is recommended Ethanol, 200 proof (EtOH) is the choice of dehydration agent for LW White. (Acetone acts as a radical scavenger in the resin system and traces of acetone left in the tissue at curing can interfere with polymerization) Conventional: 50% EtOH/50% water 15 min 70% EtOH/30% water 15 min 90% EtOH/10% water 10 min 100% EtOH 10 min 1:1 EtOH:Resin Overnight 100% Resin 2 hours 100% Resin 1 hour For Immunocytochemistry: Even though the tissue may be taken from 70% ethanol into LW White resin no special procedure is necessary. However, due to the fact that the nature of the tissue is not always consistent, we feel it is safer with at least one more change with 80% alcohol before going to resin. 50% EtOH/50% water 15 min 70% EtOH/30% water 15 min * 80% EtOH/20% Water 10 min 2:1 LR White resin to 70% EtOH 1 hour** 100% LR White resin 1 hour 100% LR White resin overnight 100% LR White resin 30 min 100% LR White resin 30 min * For enhancing contrast: add 2% PTA to this step, if post fix tissue have been treated with 1% TA. **This step is needed to avoid the tissue shrinkage due to the omitted use of osmium tetroxide during fixation. For Microwave: 50% EtOH/50% water 40 sec with temperature restriction of 37°C 70% EtOH/30% water 40 sec with temperature restriction of 37°C 90% EtOH/10% Water 40 sec with temperature restriction of 37°C 100% EtOH 40 sec with temperature restriction of 37°C 100% EtOH 40 sec with temperature restriction of 37°C 1:1 EtOH:LR White resin 15 min with temperature restriction of 45°C 100% LR White resin 15 min with temperature restriction of 45°C 100% LR White resin 15 min with temperature restriction of 45°C Polymerisation Anaerobic polymerisation is advised for curing LR White resin. Thermal (Heat) Cure: Oven temperature is set at 65°C If flat embedding is needed for easy orientation of the tissue, we recommend that you should use our PTFE flat embedding mould instead of silicone. Resin must be ‘over-filled’ in the cavities then, cut a piece of paraffin, Aclar film, or polyethylene-based film as big as the mould, and place this film right on top of the resin-filled mould to exclude oxygen. If BEEM capsules are being used, the lid is removed from the capsule. Filled the capsule and top-off with LR White resin, then use the same method as described above for flat embedding to cover the BEEM capsule. If gelatin capsules are being used, after fill the capsule with LR White resin, close the capsule and set the temperature at about 55°C (gelatin has a low melting point). Typical curing time for all the above is overnight to 48 hours, depending on conditions. Cold cure (use accelerator): Add 1 drop of the accelerator (10ml bottle) to 10ml of LR White resin and mix well. This mixture will polymerise in less than an hour. However this mixture has very poor infiltration. The resin can be cured in gelatin capsules, BEEM capsule, moulding cup tray Peel Away disposable mould. Microwave Cure: Use BEEM capsule for microwave curing. In a microwave , LR White resin will polymerise under water. Load the BEEM capsules into a Capsule Holder. Place the specimen down into the tip of the capsule. Fill the capsules with LR White resin. Put the cap on the capsules (and follow the instruction of this type of capsule holder to secure the capsule lid, so they won’t pop out during the polymerisation process). Place the whole set into the tray, fill the tray with water to cover the capsules (approx 350ml). Place it inside the microwave and start irradiating the resin as follows: 10 minutes at a temperature of 60°C* @ full power 10 minutes at a temperature of 70°C* @ full power 24 minutes at a temperature of 80°C* @ full power Let the resin block cool down completely before sectioning. *Make sure the water is at the requested temperature prior to the run Trimming and Cutting When gelatin capsules are used, an initial trimming of the block with a jewelers saw is faster and a final trim with the razor blade is then much easier. Otherwise, trimming and sectioning of a LR White block may be performed the same way as an epoxy resin block. Glass knives and or diamond knives are routinely used. Cutting speed of about 1mm per second with the thickness of 50 – 70nm is suitable. For LM, sections of 2 – 3µm is nicely obtainable (up to 15 or 20µm if required) Section Staining EM – All common stains give good results on tissue embedded in LR White resin. As an alternative to uranyl acetate, 1% phosphotungstic acid has been proven to be a good general purpose stain, both as a block stain as mentioned earlier , and as a section stain. The use of ethanol and methanol to make up stains should be avoided, because these solvents will soften the cured resin and may remove sections from supporting grids. IMMUNOCYTOCHEMISTRY – The choice of immunolocalisation technique is entirely up to the user, and PAP, hapten-anti-hapten, avidin-biotin, or gold-colloid methods may be adaptable. (See some helpful references listed below) Safety LR White is a low toxicity resin. However all acrylic resins should be considered hazardous. Work under the fume hood, along with safety gloves and goggles are always good laboratory practices. Double-gloves are sometime needed if you are in prolonged direct contact with the methacrylate resins. All chemicals including acrylic resin, should always be tightly closed after each use and stored in a safe place. If in case you have direct contact with skin or eyes, wash the affected areas with plenty of water and soap. References Fixation Ito, S. and Karnovsky, M.J. (1968) Formaldehyde/Glutaraldehyde fixative containing trinitro compounds. J. Cell Biol. 39, 168a -169a. McLean I.W. and Nakanem P.K. (1974) Peroxidase-lysin-paraformaldehyde fixative, a new fixative for immoelectron microscopy. J. Histochem, 22, 1077 – 1083 Smogyi P. and Takagi H. (1982) A note on the use of picric acid-paraformaldehyde-glutaraldehyde fixative for correlate light and electron microscopic immunocytochemistry. Neurosciences, 7, 1779-1783 Stefanini, M., De Martino, C and Zamboni, I. (1967) Fixation of ejaculated spermatozoa for electron microscopy. Nature 216, 173-174. Fixation and Embedding in L.R. White Newman, G.R. Jasani, B. and Williams, E.D. (1982) The preservation of ultrastructure and antigenicity. J. Microscopy, 127, RP5-RP6. Newman, G.R. Jasani, B. and Williams, E.D. (1982) A simple post embedding system for the rapid demonstrationot tissue antigens under the electron microscope. Histochem. J. In press. M.A. Hayat. Principles, Methods and applications, Volume 2. , ed. Academic Press, Inc. New York, pp. 38-71. J.N. Skepper and J.M. Powell. 2008a.Ultrastructural Immunochemistry (Immunostaining of London Resin(LR) White section for TEM). CSH Protocols doi:1101/pdb.top47 [Abstract/Free Full Text] Immunostaining De May, J., Moeremans, M., Geuens, G., Nuydens, R. and De Brabander, M, (1981). High resolution light and electron microscopic localization of tubulin with the IGS (Immuno gold staining) method. Cell Biol. Int. 5, 889 - 899. Hsu, S.M., Raine, L, and Fanger, H. (1981) The use of avidin-biotin-peroxidase complex (ABC) in immunoperoxidase techniques: a comparison between ABC and unlabelled antibody (PAP) procedures. J. Histochem. Cytochem. 29, 577 – 580. Jasani, B., Wynford-Thomas, D. and Williams, E.D. (1981) Use of monoclonal antihapten antibodies for immunolocalization of tissue antigens. J. Clin. Path. 34, 1000 – 1002. Larson, L. (1979) Simultaneous ultrastructural demonstration of multiple peptides in endocrine cells by a novel immunocytocheical method. Nature 282, 743 – 746. Roth, J. Bendayan, M and Orci, L. (1978) Ultrastructural localization of intracellular antigens by the use of protein A-gold complex. Stenberger, L.A. (1972) The unlabelled antibody-peroxidase and the quantitative immunoranium methods in light and electron microscopy. In ‘Techniques of Biophysical and Biochemical Morphology 1’, Rd D.R. Glick and R.M. Rosenbaum. New York, John Wiley and Sons Inc. Giberson, et al. 1997. Four-hour processing of clinical/diagnostic specimens for electron microscope . J. Vet. Diagn. Invest. 9:61-67.
LR White for Electron Microscopy
Resin embedding for light microscopy provides greatly improved cellular definition compared to paraffin embedding, and for this reason it is now widely used, particularly in diagnoses of Renal disease, Lymphomas and bone marrow trephines, as well as research. The acrylic resins currently used, however, are not suitable for E.M. and the epoxy resins used for E.M. are not easily stained for light microscopy. 'L.R. White' can be used for both purposes and a lymph node, for example, (12 x 10 x 3mm) can be processed, cut and stained for light microscopy, then the same block trimmed down, cut and stained for electron microscopy. L.R. White can also be used for the histochemical demonstration of some of the more resistant enzymes and for the immunocytochemical demonstration of intracellular immunoglobulins. For those laboratories already using an acrylic resin, e.g. HEMA or Glycol Methacrylate, no alteration need be made to the current processing schedule, but we give here a 'typical' schedule for L.R. White as guidance for its use. FIXATION: No change from normal fixation need be made if L.M. only is required from the final blocks (Neutral Buffered Formalin recommended). If E.M. is required subsequent to L.M. then we have found the use of freshly depolymerised paraformaldehyde (3-4%) in a phosphate buffer pH 7.2 with 2.5% w/v sucrose is the best compromise. Glutaraldehyde-formaldehyde mixtures may lead to very pale staining with haematoxylin and patchy eosin, whereas normal formalin fixation gives unacceptable E.M. structure. For the dual LM/EM role, osmium tetroxide should be avoided due to its effect on many LM stains but 1% phosphotungstic acid (w/v) in the first absolute ethanol step of dehydration improves electron contrast without adversely affecting most LM stains. If this does not provide adequate electron density then 'staining' of ultra thin sections can be carried out with osmium (a brief exposure to 1% aqueous osmium tetroxide or osmium tetroxide vapour on a copper grid) or lead citrate. DEHYDRATION: A graded ethanol series is the method of choice when using L.R. White. Acetone acts as a radical scavenger in the resin system and traces of acetone left in the tissue at curing can interfere with polymerisation. INFILTRATION: The extremely low viscosity of L.R. White allows the use of short infiltration times, but these will obviously depend on the size of the tissue. Infiltrated tissue will become translucent and sink to the bottom of the container. A typical dehydration and infiltration schedule for a block (12 x 10 x 3mm) on a mixer would be: Two changes 70% alcohol 30 minutes each Two changes Absolute alcohol 30 minutes each Infiltrate with L.R. White at RT 2-3 changes 60 minutes each or leave overnight. POLYMERISATION: Either heat or cold curing can be used for L.M. Cold curing gives slightly better cutting and staining qualities. When cold curing it is important to cool the moulds in a bath of cold water, during polymerisation in order to disperse the heat produced by the exothermic reaction, but it is not necessary to exclude oxygen from the surface of the curing block. Some polymerisation problems have been experienced when embedding very flat pieces of tissue which stick to the base of the embedding mould. The way to avoid this is to smear the base of the mould with accelerator before adding mixing resin, and allow the tissue to sink to the base of the mould rather than applying pressure. When thermal curing, it is important to limit the contact of oxygen with the resin while polymerisation occurs. The most convenient way of achieving this is to use gelatin capsules for small pieces of tissue. Fill up to the brim and slide the other half of the capsule on. For larger specimens the surface of the resin must be covered and one convenient method is to utilise the JB-4 type moulds, one being used as a lid for another, or to polymerise in a nitrogen environment. Polymerisation time and temperature are fundamental to the physical character of the final block, to a much greater extent than with cured epoxy systems. We strongly recommend a temperature of 60C 2C for a period of 20-24 hours. Some ovens are not capable of controlling temperature so closely and if faced with over brittle blocks this is the parameter to check. Resin may be used straight from the refrigerator and has a very low toxicity in both monomeric and polymerised states unlike epoxies (see Proc. Mic. Soc. (1981), 16, Pt.4, p. 265-271). The cold cure accelerator does have some toxic risk and contact with skin and eyes should be avoided. For cold curing the accelerator should be used at one drop per 10 ml of resin and this should cause polymerisation in 10-20 minutes. If polymerisation occurs faster than this we recommend either more careful metering of the one drop of accelerator or a higher volume of resin per drop of accelerator. CUTTING AND MOUNTING: Although it is possible to cut L.R. White on a standard microtome with a steel knife the method of choice would be to use a heavy duty motorised microtome, and glass (Ralph type) knife. L.R. White can cut as thin as 0.25 micron on some microtomes, but it is very difficult to obtain a satisfactory stain intensity with anything other than toluidine blue at this thickness, simply because there is so little tissue present in the sections. For haematoxylin and eosin staining, as well as most other routine stains, we recommend sections of 2-3 micron. It is of course possible to cut thicker (up to 15 or 20 micron) if required. Blocks can be cut dry, the sections picked up and floated out on 30-50% acetone on a hot plate @ 60-70C. To 20ml acetone add 0.5ml benzyl alcohol mix then make up to 50ml with distilled water. A section adhesive such as egg albumin, can be added to this if required. SECTION STAINING: Most routine stains give good results on tissue embedded in L.R. White resin using standard times and temperatures although it may occasionally be necessary to extend some staining times e.g. Methyl Green Pyronin. Stains made up in ethanol or methanol should be avoided as these solvents soften the resin and may remove sections from the slide. Dehydration of sections through graded alcohol after staining should also be avoided. Sections should be blotted, air dried and then mounted in a resinous mounting medium.
LR White for Light Microscopy
Using LR White for Light Microscopy Resin embedding for light microscopy provides greatly improved cellular definition compared to paraffin embedding, and for this reason is now widely used in diagnoses particularly of Renal disease, Lymphomas and bone marrow trephines as well as research. The acrylic resin currently used however are not suitable for EM and the epoxy resin used for EM are not easily stained for light microscopy. LR White however can be used for both purposes and a lymph node for example (12 x 10 x 1mm) can be processed, cut and stained for light microscopy then the same block trimmed down cut and stained for electron microscopy. LR White can also be used for the Histochemical demonstration of some of the more resistant enzymes, and for the immunocytochemical demonstration of intracellular immunoglobulins. For those laboratories already using an acrylic resin e.g. haema or glycol methacrylate no alteration need to be made to the current processing schedule, but we have laid out here a typical schedule for LR White as guidance for its use. Fixation No change from normal fixation need be made if LM only is required from the final blocks (Neutral Buffered Formalin recommended). If however EM is required subsequent to LM then we have found the use of freshly depolymerised paraformaldehyde (3-4%) in a phosphate buffer pH 7.2 with 2 1/2% w/v sucrose is the best comprise. Glutaraldehyde-formaldehyde mixtures may lead to very pale staining with haematoxylin and patchy eosin, whereas normal formalin fixation gives unacceptable EM structure. For the dual LM/EM role osmium Tetroxide should be avoided due to its effect on many LM stains but 1% Phosphotungstic acid (w/v) in the first absolute ethanol step of dehydration improves electron contrast without adversely affecting most LM stains. If this does not provide adequate electron density then “staining” of ultrathin sections can be carried out with osmium (a brief exposure to 1% aqueous osmium Tetroxide or osmium Tetroxide vapour on a copper grid) or lead citrate. Dehydration A graded ethanol series is the method of choice when using LR white. Acetone acts as a radical scavenger in the resin system and traces of acetone left in the tissue at curing can interfere with polymerisation. Infiltration The extreme low viscosity of LR White allows the use of short infiltration times, but these will obviously depend on the size of the tissue. Infiltrated tissue will become translucent and sink to the bottom of the container. A typical dehydration and infiltration schedule for a block (12 x 10 x 3mm) on a mixer would be: Two changes 70% alcohol 30 minutes each Two changes Absolute alcohol 30 minutes each Infiltrate with L.R. White at RT 2-3 changes 60 minutes each or leave overnight. Polymerisation Either heat or cold curing can be used for LM, cold curing gives slightly better cutting and staining qualities. When cold curing it is important to cool the moulds in a bath of cold water, during polymerisation, to disperse the heat produced by the exothermic reaction, but it is not necessary to exclude oxygen from the surface of the curing block. Some polymerisation problems have been experienced when embedding very flat pieces of tissue, which stick to the base of the embedding mould. The way to avoid this is to smear the base of the mould with accelerator before adding mixed resin and allow the tissue to sink to the base of the mould rather than applying pressure. When thermal curing it is important to limit the contact of oxygen with the resin while polymerisation occurs. The most convenient way of achieving this is to use gelatin capsules for small pieces of tissue. Fill up to the brim and slide the other half of the capsule on. For larger specimens the surface of the resin must be covered and one convenient method is to utilise the JB-4 type moulds, one being used as a lid for another, or to polymerise in a nitrogen environment. Polymerisation time and temperature are fundamental to the physical character of the final block, to a much greater extent than with under cured epoxy systems. We strongly recommend a temperature of 60°C ± 2°C for a period of 20-24 hours. Some ovens are not capable of controlling temperature so closely and if faced with over brittle blocks this is the first parameter to check. Resin may be used straight from the refrigerator and has a very low toxicity in both monomeric and polymerised states unlike epoxies (see Proc. Roy. Mic. Soc.(1981), 16, Pt.4, p. 265-271). The cold cure accelerator does have some toxic risk and contact with skin and eyes should be avoided. For cold curing the accelerator should be used at one drop per 10ml of resin and this should cause polymerisation in 10-20 minutes. If polymerisation occurs faster than this we recommend either more careful metering of the one-drop of accelerator or a higher volume of resin per drop of accelerator. Cutting and Mounting Although it is possible to cut LR White on a standard microtome with a steel knife the method of choice would be to use a heavy duty motorised microtome, and glass (Ralph type) knife. LR White can cut as thin as 0.25 microns on some microtomes, but is very difficult to obtain a satisfactory stain intensity with anything other than toluidine blue at this thickness, simply because there is so little tissue present in the sections. For haematoxylin and eosin staining as well as most other routine stains we recommend sections of 2-3µm. It is of course possible to cut thicker (up to 15 or 20µm) if required. Blocks can be cut dry, the sections picked up and floated out on a 30-40% acetone on a hot plate @ 60-70°C, and then allowed to dry at this temperature. For hard tissues, and blocks which contain a combination of hard and soft tissues, such as marrow trephines, the following floating out fluid is recommended, again on a hot plate @60-70°C. To 20ml acetone add 0.5ml benzyl alcohol mix then make up to 50ml with distilled water. A section adhesive such as egg albumin can be added to this if required. Section Staining Most routine stains give good results on tissue embedded in LR White resin using standard times and temperatures although it may occasionally be necessary to extend some staining times e.g. methyl green pyronin. Stains made up in ethanol or methanol should be avoided as these solvents soften the resin and may remove sections from the slide. Dehydration of sections through graded alcohols after staining should also be avoided. Sections should be blotted air dried and then mounted in resinous mounting medium.
LR White Resin for Hard Tissue
Using LR White for Hard Tissue A user note for C023, C024, C025, C026, C027. L.R. White can be used for the microtomy of decalcified bone and teeth and also for microtomy or "sawing and grinding" of undecalcified tissues. DECALCIFIED TISSUE: May be processed, cut and stained similarly to soft tissue (see Using L.R. White for Light Microscopy), except that dehydration and infiltration times may need to be extended depending on the size of tissue. It is also recommended that bone be "de‐fatted" to improve the penetration of resin into marrow cavities. This can be achieved by using chloroform after dehydration and returning to absolute alcohol to remove the chloroform before infiltrating with resin and polymerising. UNDECALCIFIED TISSUE: Dehydration and infiltration times will vary depending on size and density of tissue. Those laboratories using Methyl or Butyl methacrylate at present can use similar dehydration times, but infiltration will probably be shortened due to the low viscosity of the resin. DEHYDRATION: A graded series of alcohol should be used for dehydration of tissue, and when processing bone "de‐fatting" is recommended to improve the penetration of resin into marrow cavities. This can be done using chloroform for the same length of time that would be necessary to clear the tissue. The bone should then be taken back to absolute alcohol and given sufficient changes to remove the chloroform before infiltration with L.R. White (Hard Grade). INFILTRATION: Several changes of resin will be necessary and impregnation under vacuum is recommended. POLYMERISATION: The tissue can be heat or accelerator cured after embedding in strong plastic moulds, such as JB4 or Peel‐a‐way type, or aluminium foil dishes. When heat curing the moulds should first be filled with resin then the tissue added and oriented. Polymerisation will occur in 18‐24 hours at 60-62ºC. The surface of the block exposed to oxygen may remain slightly sticky, but this will not affect the cutting quality of the face of the block. Some ovens are not capable of controlling temperature so closely and if faced with overly brittle blocks, this is the first parameter to check. When accelerator or 'cold' curing the moulds should be placed in a bath of ice‐cold water to disperse the heat produced during the exothermic polymerisation. The base of the moulds should be smeared with accelerator using a cotton‐wool bud or swab, the accelerator is then added to the resin, 1 drop per 10ml resin, and thoroughly mixed before pouring into the mould. The tissue is then placed into the mould and oriented. Polymerisation should occur in 10‐20 minutes. If it occurs faster than this, we recommend either more careful metering of the one drop of accelerator or a higher volume of resin per drop of accelerator. N.B. the accelerator does have some toxic risk and contact with skin and eyes should be avoided. CUTTING AND MOUNTING: Bone marrow trephines and small pieces of cancellous bone may be cut using a motorised heavy duty microtome, but larger pieces of cancellous bone, cortical bone and teeth offer too much resistance to the microtome knife and preparations of this material must be prepared by sawing and grinding. MICROTOMY: Sections can be cut, using Ralph type glass knives for trephines or a tungsten carbide knife for larger pieces of cancellous bone, from 2‐10 micron. Blocks can be cut dry, the sections picked up and floated out on a hot plate at 60‐70ºC using the following solution: to 20 mL acetone add 0.5ml benzyl alcohol mix then make up to 50ml with distilled water. A section adhesive such as egg albumin can be added to this if required. Sections should be allowed to dry on the hot plate for at least 30 minutes before staining. STAINING AND GRINDING: Thick slices 150‐200 micron can be cut using a milling machine and then ground to the required thickness, usually 20 micron for staining or 70 micron for microradiography; the section is inclined to fragment if grinding is continued much below 20 micron. Using the newer types of saw microtome, such as the Leitz 1600, which has a diamond‐coated‐internal‐hole‐saw, sections can actually be cut at 20 micron and no further grinding is necessary. SECTION STAINING: Sections of material embedded in L.R. White are stained "free floating". Times of staining are usually longer than those for paraffin sections, and dehydration through alcohol should be avoided. A recommended schedule for Haematoxylin and Eosin staining is as follows: Remove Ca deposits, which would otherwise interfere with the staining, by treating with Kristensene's Decalcifying Solution for about 15 minutes. Wash in running tap water for a few minutes to remove the formic acid from the tissue. Transfer the section to several changes of distilled water, a few minutes each, and then into a 0.5% w/v solution of periodic acid in distilled water where it is left for 5 minutes. Wash the section to several changes of distilled water and stain with Harris Haematoxylin for about 1 hour. Transfer the section, after a short time in distilled water, into running tap water to "blue" the Haematoxylin stained tissue. Check the Haematoxylin stain; if the tissue should be overstained or the surface of the resin has become stained with Haematoxylin, this may be corrected by a short differentiation in acid alcohol (0.5% HCl in 70% ethanol) and "reblueing" of the stained tissue. Rinse the section in distilled water and counterstain it, using a 5% solution of Eosin Y in distilled water; leave the section in the Eosin stain for 30 minutes to 1 hour, wash briefly in running water and check the staining of the tissue. Nuclei and haematoxophilic elements should be bright blue, cytoplastic structures in various shades of red‐pink. Rinse the section in distilled water, blot dry with filter paper and either clear briefly in xylene and mount in DPX, or mount directly in L.R. White resin by adding a drop of accelerator to 1ml of resin.
LR White Resin for Immunocytochemistry
A user note for C023, C024, C025, C026, C027. Sections from L.R. White embedded tissue have been used successfully for immunocytochemistry at both the light microscope and electron microscope levels. This demonstrates quite clearly that the visualisation steps of the immunocytochemical procedure will penetrate the resin and react with the tissue antigens if they have been preserved in the tissue. As with all immuno‐localisations, the key factor is whether or not the tissue antigen has survived fixation, processing and embedding in such a form as to be recognisable to the specific antibody. This is difficult to predict with certainly, but some antigens have been shown to be highly resistant whilst others are fickle even in unfixed frozen sections of antisera Much interest has centered on using immunocytochemistry to detect protein hormones and the various classes of immunoglobulin and generally these classes of antigen have proved resistant to alteration both in processing to paraffin‐wax and to L.R. White. It is the special hydrophilic nature of L.R. White which allows immunochemicals to permeate the supporting resin and reach its sites of binding and no resin pretreatment is necessary, or indeed possible, to facilitate this penetration. We have been successful with L.R. White blocks only when they have been thermally cured, probably because when accelerator‐curing the resin the exotherm produced is sufficient to damage the integrity of the tissue antigen. Some workers have also reported that a slight under‐cure of L.R. White, say at 55ºC for 20‐24 hours, aids subsequent penetration, but we have obtained good results without deviating from the standard polymerisation schedule. If the particular antigen under consideration has already been localised in paraffin‐wax sections then a trusted fixation regime will be established and should be adhered to. For those approaching the problem for the first time there is an extensive bibliography available regarding fixation for immunocytochemistry, much of it contradictory, and a reference list is provided as some guidance. Rules of thumb seem to be to avoid glutaraldehyde and perhaps use an acid rather than a neutral fixative, but there are many conflicting and strongly held views on the topic. Similarly, the need to enzyme digest sections prior to reaction is fraught with controversy and may indeed be linked to the fixation regime chosen. We have used both protease type VII and trypsin type II to good effect on our neutral buffered formalin fixed material. If frozen, dewaxed or etched epoxy resin sections are used for immuno‐staining, the tissue is not surrounded by a supporting matrix when they are being reacted. When using L.R. White sections the resin is still intact and therefore diffusion to the sites of reaction must occur prior to reaction of antisera with antigen. For this reason we have found it necessary to use antisera at approximately ten times the concentration that would work on de‐waxed sections. The exact titre of each antibody will depend upon its source and how well it has been stored, but we have used many commercial anti‐immunoglobulins at about a 1 in 10 dilution. For the same reason the antibody stages of the reaction often benefit from a longer incubation time. Up to 2 hours at room temperature or overnight at 4ºC in a moist chamber may be used. Various immunoperoxidase techniques have given results on L.R. White tissue sections including the peroxidase‐antiperoxidase complex method (PAP) (Sternberger, 1970), the hapten sandwich technique (Jasani et al, 1981), and the indirect peroxidase method. The Avidin‐biotin‐peroxidase complex method of Hsu has not been successful in our hands with L.R. White embedding material, probably due to the molecular size of Avidin. As fairly strong antibody concentrations are required, a highly sensitive method of detection is to be preferred and for this reason the PAP or hapten sandwich techniques are more suitable than a two layer indirect peroxidase reaction. Visualisation of the bound peroxidase is achieved with the diaminobenzidine‐peroxide reaction as described by Graham and Karnovsky (1966). Any technique where the sections are subjected to hydrogen peroxide solutions twice during staining is likely to tend to lift sections from the slides. We have found that lysine (MW 350,000) is an excellent adhesive for immunocytochemical work, and also care should be taken to dry sections onto slides very thoroughly in an oven rather than a hot plate at 60 degrees C for two hours. This step should not have any effect on the antigenicity of the tissue as it will already have spent 20‐24 hours at 60ºC during polymerisation. It is clear that no 'standard' immunohistochemical staining regime can be cited as there are so many variables, but a 'typical' regime is described below for general guidance. Pap Procedure for L.R. White Sections (3 micron) Block Endogenous Peroxidase with 1% Phenylhydrazine Hydrochloride in P.B.S. (optional) 30 minutes Wash in P.B.S 1 x 5 min, and 2 x 5 mins at 37ºC 0.1% Trypson in 0.1% CaCl2 (aqueous) 20 minutes Wash in ice cold distilled water 10 minute 2% Goat serum in PBS 20 minutes 1° Antibody (approx 1:10 dilution) 2 hrs at 37ºC or overnight at 4ºC (usually Daco or Nordic) Wash in P.B.S. 10 minutes Goat anti rabbit antibody (approx 1:20 dilution) 2 hrs at 37ºC or overnight at 4ºC Wash in PBS 10 minutes PAP at 1:10 dilution (Dako) 2 hrs at 37ºC or overnight at 4ºC Wash in PBS 10 minutes Wash in Tris HCL pH 7.6 10 minutes DAB/H2O2 (Graham and Karnovsky) 15 minutes Wash in distilled water. Counterstain as required NOTE: FOR IMMUNOCYTOCHEMISTRY L.R.WHITE MUST BE THERMALLY CURED AND NOT ACCELERATOR CURED References: CURRAN, R.C.and GREGORY, J., The unmasking of antigens in paraffin sections by Trypsin. Experientia, 33; 1400 (1977) ELIAS, J.M., Principles and Techniques in Diagnostic Histopathology. Published by Noyes Publications New Jersey, USA (1982) ISBN‐0‐8155‐0903‐0. GRAHAM, R.C.and KARNOVSKY, M.J., The early stages of absorption of injected horseradish peroxidase in the procimal tubules of mouse kidney: ultrastructural cytochemisty by a new technique. J. Histochem. Cytochem, 14, 291 (1966) JASANI, B., WYNFORD‐THOMAS, D.,and WILLIAMS, E.D., Use of monoclonal anti‐hapten antibodies for immunolocalization of tissue antigens. J. Clin. Path. 34, 1000 (1981). NEWMAN, G.R., JASANI, B., and WILLIAMS, E.D., The preservation of ultrastructure and antigenicity, J. Microscopy, 127, RP5‐RP6 (1982). STERNBERGER, L.A., HARDY, P.H. Jr., CUCULIS, J.J. and MEYER, H.G., The unlabelled antibody enzyme method of immunohistochemistry. Preparation and properties of soluble antigen‐antibody complex (horseradish peroxidase‐antihorseradish peroxidase) and its use in identification of spirochetes. J. Histochem. Cytochem., 18, 315 (1970)
LR White Resin, Uncatalysed
C023 & C024 LR White resin is supplied uncatalysed; with the catalyst (Benzoyl Peroxide) supplied 9.9g in a separate tube. LR White (catalysed) should be kept at 4°C or lower. In our experience, the shelf-life of LR White is a minimum of one year @ 4°C, and up to 15 months if stored as low as -80°C. If the resin is frozen, allow it to thaw and reach room temperature prior to use. If the viscosity of the LR White resin changes (becomes thicker), stop and do not use it. LR White can be polymerised by four different methods: Heat: 60 – 65°C UV irradiation (365nm wavelength) Chemical with accelerator Microwave Mixing LR White Resin with Catalyst (Benzoyl Peroxide): All of the catalyst should be added to resin at room temperature and the resin must be shaken vigorously for a couple of minutes. There have been incidents where a whole bottle hardened. It must be understood that much of the catalyst will dissolve within an hour, but if after initial shaking the bottle is left without occasional inverting, and then the catalyst settles and initiates an exothermic reaction. The heat further accelerates the reaction and an hour later the whole bottle may be very hot and solid. To avoid this: Manually invert the bottle frequently every few minutes at first, increasing to every ten minutes. Using a magnetic stirrer, but never use heat. Ensure that the magnet is stirring and not just sitting at the edge of the bottle. If a slow rotator or shaker is available (not in a hot room) squeeze most of the air from the bottle after the catalyst is added, lay the bottle on the side, then rotate for a couple of hours. Refrigerate the bottle after about three hours (two hours is sufficient if it had been mechanically agitated). Usually it takes 12 hours at room temperature and 12 hours in the refrigerator for catalyzed LR White resin to take effect prior to use. Do not attempt to heat the resin in order to speed the dissolution of the catalyst. Once mixed and fully dissolved the resin must be stored at 4°C to maintain its shelf life. Freshly catalysed and thoroughly oxygenated resin may take a little longer than normal to polymerise. Following the addition of the catalyst, a test aliquot should be polymerised at 60°C for twenty-four hours as a quality control measure. We do not advise catalysing less than 500g (one full bottle) of LR White resin as it can be difficult to accurately measure smaller quantities of the reagents. LR White resin with insufficient catalyst will normally polymerise by thermal curing eventually, though curing times may be protracted. Uncatalysed LR White with accelerator only added may only cure to a gel. References Fixation Ito, S. and Karnovsky, M.J. (1968) Formaldehyde/Glutaraldehyde fixative containing trinitro compounds. J. Cell Biol. 39, 168a :169a. McLean I.W. and Nakanem P.K. (1974) Peroxidaselysin paraformaldehyde fixative, a new fixative for immunoelectron microscopy. J. Histochem, 22, 1077: 1083 Smogyi P. and Takagi H. (1982) A note on the use of picric acid/paraformaldehyde/glutaraldehyde fixative for correlate light and electron microscopic immunocytochemistry. Neurosciences, 7, 1779:1783 Stefanini, M., De Martino, C and Zamboni, I. (1967) Fixation of ejaculated spermatozoa for electron microscopy. Nature 216, 173:174. Fixation and Embedding in L.R. White Newman, G.R. Jasani, B. and Williams, E.D. (1982) The preservation of ultrastructure and antigenicity. J. Microscopy, 127, RP5:RP6. Newman, G.R. Jasani, B. and Williams, E.D. (1982) A simple post embedding system for the rapid demonstration of tissue antigens under the electron microscope. Histochem. J. In press. M.A. Hayat. Principles, Methods and applications, Volume 2. , ed. Academic Press, Inc. New York, pp. 38:71. J.N. Skepper and J.M. Powell. 2008a.Ultrastructural Immunochemistry (Immunostaining of London Resin(LR) White section for TEM). CSH Protocols doi:1101/pdb.top47 [Abstract/Free Full Text] Immunostaining De May, J., Moeremans, M., Geuens, G., Nuydens, R. and De Brabander, M, (1981). High resolution light and electron microscopic localization of tubulin with the IGS (Immuno gold staining) method. Cell Biol. Int. 5, 889 :899. Hsu, S.M., Raine, L, and Fanger, H. (1981) The use of avidin-biotin-peroxidase complex (ABC) in immunoperoxidase techniques: a comparison between ABC and unlabelled antibody (PAP) procedures. J. Histochem. Cytochem. 29, 577 : 580.
Meltmount™ Quick-Stick™
I011-1539, I011-1582, I011-1605, I011-1662, I011-1680 & I011-1704 Meltmount Quick-Stick For a number of years Cargille Laboratories has been using the Meltmount Quick-Stick for the preparation of Cargille-Allen reference set slides. Notes: Do all work in a well ventilated area Meltmount is a thermoplastic: it is fluid when heated and functionally a solid at room temperature; the appearance of the prepared slide will remain unchanged after the slide is returned to room temperature. The most common problem encountered is: the inclusion of bubbles in the mount. The procedure described here should reduce or eliminate this problem. Meltmount Quick-Stick is conveniently enclosed in a Teflon® tube that can be cut back to expose the Meltmount so that it can be applied directly to a heated slide, making slide preparation quick and neat. Meltmount is a Thermal Plastic Material. This means its viscosity is dependent on temperature (inversely dependent). As the temperature increases the viscosity decreases. There is no sharp melting point. Being thermal plastic, it is capable of "cold flow". This means the Meltmount, the specimen, the slide, and the cover slip can all move independently of each other given a mix of time, temperature, and lateral pressure or gravity. Storage of prepared slide: treat them as the valuable items they are. Store: Flat, cover slip on top In the dark Away from dust and fumes At 15° to 30°C. Meltmount is meant to be thermally reversible, don't allow this to happen inadvertently by storing or transporting prepared slide above 29°C. The Procedure for Using Meltmount™ Quick-Stick™ Adjust hot plate for medium heat (ideally 60°C to 70°C). Remove the tall cap bearing the label. When using Meltmount Quick-Stick I011-1582, I011-1662, I011-1680 and I011-1704 (but not when using I011-1539), firmly roll the end to be used between your thumb and index finger or on a hard surface such as a lab table top; this will break the adhesion of the Meltmount to the Teflon tube. Use a single edged razor blade to cut off and remove the Teflon to expose ¼ to ½ inch of Meltmount. Put a trial slide on the hot plate and apply Meltmount to its center. The melted Meltmount on the slide should be thin and watery but not smoking. If not, adjust the temperature up to make thinner or down to avoid smoking (60°C to 70°C is ideal). Put a slide on the hot plate with about ¼ of the slide off the hot plate's surface where it will remain cool so it can be handled. Use the Quick-Stick to apply a 2 cm patch of Meltmount to the centre of the slide. Put the specimen in the centre of the patch of the Meltmount, you may want to remove the slide from the hot plate to do this, and then return it to the hot plate. Do the following as quickly as possible Place the cover glass completely on the hot plate surface. Apply a 2 cm size patch of Meltmount to the centre of the coverglass. Raise the Quick-Stick from the hot plate WITH the cover glass attached to the Quick-Stick. Remove the cover glass from the Quick-Stick, invert and drop (Meltmount side down) onto the centre of the slide. Use a pencil eraser to centre cover glass while pressing to remove bubbles. Remove mounted slide from the hot plate. Note: Heat Sensitive Specimens may be altered by this method, in which case, mounts can be made without heat by The Pressure Method described later in these instructions. If the area beneath the cover glass is not completely filled with Meltmount, return the slide to the hot plate and add more Meltmount by touching the Quick-Stick to the edge of the cover glass. Then remove from the heat. Optional: The cleaning of the mounted slide and removal of excess Meltmount can be done as follows:When the slide has cooled to room temperature, scrape off excess Meltmount using as a tool a single edge razor blade or dental scraping tool. The tool can be dipped frequently in cold water to keep it cool and avoid sticking. Soak the slide in a tray of Windex® window cleaner. While immersed, clean around the edge of the cover glass using a sponge tipped swab (a cotton tipped swab will leave fibres behind). Remove the slide from the tray and do the final cleaning with a lint free tissue. The Pressure Method Note: This method is useful for making permanent slides in the field, or for making permanent slides of heat sensitive specimens. Turn on hot plate to medium heat (ideally 60°C to 70°C). Place clean slide on hot plate with ¼ of the slide off the hot plate so that this portion will remain cool so that you can pick it up with your fingers. With the Quick-Stick apply a patch of Meltmount that is the size of a coverglass. Remove slide from hot plate and cool to room temperature. Store slides with Meltmount in a slide box, with slides held flat. Applying the specimen can be done in many ways such as: Drop specimen onto the patch of Meltmount Place the Meltmount patch on the slide directly against the specimen. Transfer specimen to sticky tape, then transfer from tape to the Meltmount patch on the slide by running your fingernail over the back of the tape. The cover glass can be used either with or without first applying to it a layer of Meltmount. The Meltmount can be applied to one side of the cover glass as it sits on the hot plate. When the cover glass has been removed and has cooled it can be placed over the patch of Meltmount containing the specimen on the slide. In the field it may be more convenient to place a cover glass WITHOUT Meltmount on top of the Meltmount patch containing the specimen on the slide. Press the cover glass and slide together using thumb and forefinger, taking care not to crack the cover glass. If desired, the finished slide can be improved by one of the following methods: Apply pressure of thumb and forefinger for a longer time. Heat briefly on a hot plate. Place a waxed paper covered weight, such as a book, on top of the mounted slide overnight.
Methacrylate Embedding Media
Methacrylates embedding media consists of a mixture of n-Butyl and Methyl methacrylates with benzoyl peroxide as the catalyst. (Benzoyl peroxide is supplied as a paste, a mixture of benzoyl peroxide powder with dibutyl phthalate 1:1 w/v.) The desired hardness of the block can be obtained by adjusting the relative proportion of n-butyl and methyl methacrylates; a higher proportion of methyl methacrylates will produce harder blocks. Relatively soft blocks are good for soft tissues, such as pancreas, while harder blocks are suited for tough tissues such as tendon, muscle, and plant. The hardness of the blocks is obtained by mixing n-butyl and methyl methacrylates in the ratio of 4:1; 3:1; 3:2...(v/v) respectively. The ratio of 4:1 is the most commonly used. The addition of initiator, benzoyl peroxide paste, is at a ratio of 1.5-2% of the total volume of the mixture. Methacrylates embedding may be used for embedding mineralised or unmineralised specimens. Because of the large shrinkage that occurs during polymerisation of methacrylates, direct embedding is not recommended for most biological tissues, and it is therefore partially polymerised methacrylates (or pre-polymerised) which is required. Using Methacrylates Embedding Media 1. Preparation of Pre-polymerised MBM: It is not necessary to remove polymerisation inhibitors from the methacrylates. The mixture of methyl and butyl methacrylates plus catalyst (MBM) is prepared by mixing n-butyl and methyl methacrylates per the ratio that you chose with the addition of 1.5-2% benzoyl peroxide paste. Mix well, then place the mixture in a water bath at 65°C until the temperature of the solution slightly exceeds 65°C. The reaction temperature of the solution should be maintained between 65 and 70°C until the viscosity of the liquid resembles that of room temperature glycerine (about 30-40 minutes). The reaction temperature is not allowed to exceed 70°C. The solution then is cooled in an ice bath and stored in a refrigerator. This pre-polymerised stock solution is stable for about one year when kept refrigerated. Preparation of Methacrylates Catalyst Stock Solution: The methacrylates catalyst stock solution consists of 2 g benzoyl peroxide paste and 18 g methyl methacrylates monomer. Mix well. The catalyst stock solution is then stored under refrigeration. Embedding mixture: Every 1 ml of catalyst stock solution is mixed well with 5 ml of prepolymerised MBM. Embedding Schedules: Methacrylates are readily soluble in ethanol and acetone and no intermediate solvent is required. The standard embedding schedule is: Ethanol/MBM 1 hour MBM 1 hour MBM 1 hour Place specimens in gelatine or polyethylene embedding capsules and fill the capsules with the embedding mixture. Close the capsule with their caps. Polymerisation takes place over night at 60-65°C or 24-48 hours under UV light.? Sectioning and Mounting: The sections should be cut on a dry knife (Diamond or Glass Knife) 1 to 3 microns in thickness. Sections can be picked up and floated on a drop of water on a glass slide that has been treated with a thin coating of an albumin/glycerine (1:1, v/v). The slide is then placed on a hot plate and heated for a few seconds to dry the water drop and to adhere the section to slide. Prior to image analysis or staining, contrast is enhanced by immersing the slide in 2-butanone to remove the embedding medium. References: A.M. Glauert (1975). Practical Methods in Electron Microscopy. North-Holland Publishing Company. M.A. Hayat (1989). Principal and Technique of Electron Microscopy. 3rd Edition, CRC Press, Inc. E.K. Boylston et al., (1995). A quick Embedding Method for Light Microscopy and Image Analysis of Cotton Fibers. Biotech. & Histochem. pp. 24-27
Osmium Tetroxide
C010 & C012 This chemical is the most commonly used E.M. fixative for biological specimens. As it is a very toxic substance, safe handling procedures and adequate facilities should be available to all users. IMPORTANT: Basic safety considerations before opening product Work with Osmium Tetroxide only under a fume hood to prevent inhalation and contact by OsO4 vapour, particularly of the eyes which are very sensitive to exposure. Prevent contact with skin by wearing gloves. Never pipette osmium by mouth! Preparation of a stock solution 1. Commercial OsO4 is supplied in sealed glass ampoules. Handle osmium only under a fume hood, with the sash lowered to protect the operator's face and improve the draw. Plastic or rubber gloves are recommended. 2. Remove label and soak the vial in water to remove any glue. The solution is conveniently prepared in a wide mouth, preferably brown, glass stoppered bottle. 3. Place the open bottle on a sheet of paper and open the vial above the paper also. Unscored vials may be scored with a diamond pen or a triangular file and then opened. Scored vials may be opened with a pull and bend motion with both thumbs behind the score line. Both parts of the opened vial are dropped into the jar. Alternatively, a hot glass rod, pressed against the score-line, produces a crack around the ampoule. Any crystals spilt onto the paper should be added to the bottle. 4. To make a 2% solution of OsO4: add 50ml of distilled water to 1g of Osmium Tetroxide crystals. Crystals dissolve within about two days at refrigerator temperature. Five minutes of ultrasound breaks up crystals and aids solution. Preparation of fixative: The aqueous osmium stock solution may be mixed 1:1 with double strength buffer. Alternatively, OsO4 may be prepared complete with the desired buffer. However, some chemicals (e.g. sucrose) cause rapid reduction. Preparation of double strength aqueous osmium and double strength buffer solutions gives greater flexibility for the selection of buffers, pH and other variables. For special requirements OsO4can be made up in water. It also dissolves rapidly in organic solvents. Storage: Osmium tetroxide ampoules should be stored in a cool, dark place. OsO4 solutions are quickly reduced at room temperature and will turn from clear/yellowish to brown and then black. Brown or black solutions are exhausted and should not be used as a fixative. Stock of a working solution should be refrigerated. Alternatively, small volumes may be dispensed in glass vials (check that these will not crack with water during freezing) and store in a freezer. Physical Properties: Osmium Tetroxide (Osmic Acid) is a translucent, yellowish, crystalline substance. The melting point is 40°C. It has a high vapour pressure and pungent odour. The chemical is a powerful oxidising agent and is rapidly reduced at room temperature. Solubility in water is poor. Threshold Limit Value as Osmium in the work place is 0.0002ppm; this concentration is too low for our sense of smell. Excessive exposure results in lung congestion, skin and eye damage. Disposal: The small quantities used in laboratories can be flushed down the sink in a fume hood. It is dangerous to stockpile old OsO4 solutions in the refrigerator. Spill control: A major spill could require the evacuation of a building, whereas a spill inside a fume hood could be almost inconsequential. When heaped over spilled Osmium Tetroxide, full-cream powdered milk will exhaust crystals, solution and vapour. Before re-entering a room after a spillage, ventilate the room well; even for smaller spills, gas-tight eye goggles are needed. Neutralisation: A 2% solution of Osmium Tetroxide can be fully neutralized by twice the volume of oil (corn oil is preferred because of its high percentage of unsaturated bonds).1 That is to say, for every 10ml of 2% Osmium Solution, 20ml of corn oil is required. First Aid: Eyes: Irrigate with water for at least 15 minutes. Seek medical advice. Skin: Remove contaminated clothing and wash affected skin with plenty of water. Inhalation: Remove from contaminated area, loosen clothing and administer oxygen if necessary. Seek medical advice. Ingestion: Drink large quantities of milk or water. Do not induce vomiting. Seek medical advice.
Para-Tissuer Pen
ID600 The Para-Tissuer provides a thin coating of the adhesive on the glass slide. A sticky coating will help prevent paraffin-embedded sections from falling off, moving, or wrinkling on the slide. The Para-Tissuer coating will flatten the sections even in room temperature water and reduce the processing time. The pen can be effective with PAP/PAAP, ABC, LAB-SA and immunofluorescent methods. A new addition to the popular PAP Pen family of microscopy accessories. DIRECTIONS FOR USE: Coat an area on the uncoated slide larger than the section to be mounted. (Section may fall off if the slide is pre-treated). Let dry at room temperature for several minutes before mounting the section. Use only dry slides for the routine procedure. Use dryer if the environment is colder and high in humidity. The formulation is stable up to 120°C. Keep the tip of the pen clean to avoid contamination. Impurity may alter the result of the staining. Cap the pen tightly after use. Store with capped side down. The Pen should be discarded one year after opening.
PEEK Polish
EMS71833-10, 71833-11, 71833-12, 71833-13, 71833-14 PEEK Polish is a non-corrosive, non-irritant, non-oxidant product. It can be used in a variety of areas including the household, marine industries, and aeronautical industries and more. Not only does PEEK Polish clean surfaces, but it protects them as well. What Does PEEK Polish Clean and Protect? PEEK Polish cleans, polishes, and protects the following: Brass, silver, copper, gold, chrome, stainless steel, bronze, pewter, fibreglass, plexiglass, acrylic, aluminium, ceramic tile, marble, melamine, formica, arhorite, sterling silver, and more. What Does PEEK Polish Remove? PEEK Polish removes the following: Oxidation, tarnish, lime deposits, corrosion, crayon, grease, salt water stains, leaf marks, tree sap, bugs, tar, black marker, finger prints, graffiti, minor rust, ink, food stains, blood stains, smoke damage, water marks, wax build-up, gun powder residue, heat discolouration, and more. Tips and Directions Apply PEEK Polish sparingly. Do not let it dry, Rub onto surface to be cleaned using a dry or damp cloth. For large areas, use a damp cloth. On large surface areas, PEEK Polish may also be applied using a low-speed buffer. On intricate surfaces, PEEK Polish may be applied using an old toothbrush wrapped in a dry or wet polishing cloth. On minor rust or stubborn deposits, PEEK Polish may be applied with a nylon scoring pad or bronze wool. Using a circular motion, polish the surface until the polish has settled completely into the surface.
Photo-Flo 200 Solution
EMS74257 Description: Concentrated wetting agent used as a final step in processing to inhibit the formation of water spots or streaks on films and plates. Generally used on a silver-based black and white films and plates. Not intended for us a as a replacement for stabilisers or final rinses in colour film processes. Also used as a surfactant to aid in the smooth application of water-based retouching materials to processed films, plates and papers, both black and white and colour. Available in three different concentrations to accommodate all users, from darkroom hobbyists to large scale professional labs. When mixed according to directions, all three concentrations result in equivalent working solutions. Recommended for us in water only. Not intended to be added to processing solutions. Warning: Observe precautionary information on containers and in Material Safety Data Sheets! Photo Flo Solution is an eye irritant. Wear adequate eye protection when mixing and using this product. Avoid contact with clothing or prolonged contact with skin
PIPES Buffer
1,4 Piperazine bis (2‐ethanosulfonic acid) Useful pH range: 6.4 ‐ 7.2 Reference: Good, N.E., et al., J. Biochem. 5, 467 ‐ 1966 Baur, P.S., et al., J. of Microscopy, 109, 315 ‐ 1977 Introduction PIPES is a hydrogen‐ion organic‐based buffer, used in the preparation of a variety of mammalian tissues for EM. It produces excellent results in the preservation of ultrastructural details, especially when long fixation is required. This buffer does not appear to contribute extraneous anions or cations to the tissue, thus permitting elemental determinations by means of energy dispersive X‐Ray analysis. Baur, et al., have demonstrated that the ultrastructural aspects of various tissues (human skin, human hypertrophic scars, mollusc neuronal tissue, oyster gill, hamster ovary, cell in culture, rat skin, lung and skeletal muscle, and canine cardiac muscle) fixed with NaOH‐PIPES‐bufferedglutaraldehyde solution were of superior quality with more detail between organelles. The cytoplasm is more homogeneous in consistency, mitochondria is very electron‐dense, microfilaments and microtubules are more commonly observed in PIPES tissue and to a lesser degree in phosphate‐treated cells. The following procedure is recommended Fixation for 24 to 48 hours in a 3% glutaraldehyde, 0.1M PIPES solution buffered with NaOH to pH 7.6. Rinse the PIPES buffer for 20 minutes, using three changes. Post fixation is with 1% 0s04 with 0.1M PIPES buffer for 1 ‐ 2 hours. Other Useful Chemicals for Electron Microscopy Sciences PIPES ‐ C8H18N206S2 M.W. 353.3 ‐ C022 8% solution ‐ Glutaraldehyde EM grade ‐ 10mL ampoules 10% solution ‐ Glutaraldehyde EM grade ‐ 10mL ampoules 25% solution ‐ Glutaraldehyde EM grade ‐ 10mL ampoules ‐ C001 50% solution ‐ Glutaraldehyde EM grade ‐ 10mL ampoules ‐ C003 Osmium tetroxide, 0s04 ‐ one gram ampoule ‐? C010 Osmium tetroxide ‐ 4% aqueous solution ‐ 5mL ampoules Osmium tetroxide ‐ 4% aqueous solution ‐ 10mL ampoules ‐? C011 Sodium hydroxide pellets ‐ 100 grams ‐ C201
Procure 812 Embedding Kit
PROCURE 812 is our replacement for EPON 812, the most widely used embedding resin for electron microscopy, which was discontinued in 1978. PROCURE 812 provides the same excellent preservation and cutting qualities as EPON 812, and may be substituted in all similar formulations. Luft (1961) established EPON 812 as a reliable embedding medium, excellent both for plant and animal tissue. The same advantages offered by EPON 812, are offered by PROCURE 812: rapid penetration, greater contrast, easy sectioning, stability under the electron beam, satisfactory staining of most thick sections for light microscopy and thin sections for electron microscopy. Recommended Procedure: Fixation: Tissues can be fixed in a wide range of fixatives. One of the more commonly used fixatives is an aldehyde (i.e.: glutaraldehyde) followed by osmium tetroxide. Dehydration: There are many different dehydration schedules that can be followed. A typical one is as follows: 70% Ethanol for 10 minutes 100% Ethanol for 10 minutes 100% Ethanol for 15 minutes 100% Propylene Oxide for 15 minutes 100% Propylene Oxide for 15 minutes **NOTE: Longer times may be required for some samples. MIXING INSTRUCTIONS: Mixture A: Small Amount Medium Amount Large Amount PROCURE 812 DDSA 5ml 8ml 20ml 31ml 62ml 100ml Mixture B: PROCURE 812 NMA 8ml 7ml 20ml 17ml 100ml 90ml Final Embedding Mixture: Mixture A: 13ml 51ml 162ml Mixture B: 15ml 37ml 190ml DMP-30* 0.42-0.56 ml 1.3-1.7ml 5.3-7.0ml * For better penetration and stability, BDMA is recommended in place of the DMP-30. The quantity of BDMA which is required is 2.5-3% while DMP-30 is 1.5-2%. It is much simpler when mixing PROCURE 812 to use a one-step single mix formula. The following formulations may be used depending on the desired hardness of the block: Soft Medium Hard PROCURE 812 20ml 20ml 20ml DDSA 22ml 16ml 9ml NMA 5ml 8ml 12ml DMP-30 0.70-.94ml or 1.18-1.4 ml (BDMA) 0.66-.88ml or 1.1-1.3ml (BDMA) 0.62-.82ml or 1.0-1.2ml (BDMA) Slight variations of the accelerator (DMP-30 or BDMA) will drastically affect the colour and brittleness of the block. Prior to measuring and mixing, the resin and the anhydride should be warmed (60°C) to reduce their viscosity. Immediately before use, the two mixtures (A&B) are blended, and the accelerator added in the above-mentioned proportion. Thorough mixing is imperative to be able to achieve uniform blocks. While preparing PROCURE 812, the hardness of the block can be varied to suit various sectioning conditions depending on the ratio of mixture A and mixture B in the final embedding mixture. An increase in the proportion of mixture B will make the block harder. A mixture of 1:1 has proven most successful for general use. All components of the kit should be kept at room temperature in tight stoppered bottles. Although the mixture can be stored for up to 6 months at 4°C it is highly recommended that freshly prepared embedding medium always be used. If you choose to store the mixture you should warm it thoroughly prior to adding the accelerator. INFILTRATION: It is recommended that for all of the infiltration steps a specimen rotator be used. Drain the tissue of most of the propylene oxide, leaving a little so the tissue does not dry out. Replace the solvent with a 1:1 solution of propylene oxide embedding medium and allow it to stand for at least 1 hour at room temperature. A second change of 2:1 embedding medium to propylene oxide at room temperature overnight is recommended. Remove the mixture, replace it with 100% embedding medium and leave for 30 minutes-2 hours at room temperature. EMBEDDING: This may be done in embedding capsules or a flat embedding mould. Transfer each sample to a dry capsule or mould and fill the mould with embedding medium. Cure the medium in an oven at 60°C for 24 hours. Blocks can be trimmed and sectioned after the blocks return to room temperature. REFERENCES: Luft, J.H.(1961), J. Biophys. Biochem. Cytol. 9, 409.
Procure-Araldite Embedding Kit
Procure-Araldite recommended procedure and information. The combination of different embedding resins is popular because these kits blend the best qualities of each individual resin into one. In 1964, Mollenhauer developed an Araldite-Epon (PROCURE-812) mixture for embedding plant tissue, with particular interest in preserving the plant cell wall. It has also shown to be successful with animal tissue. Sectioning with this mixture of resins has proven to be easier than with the individual resins and the thermal stability of Araldite and the image contrast of PROCURE has made this a very successful blend. Recommended Procedure: Fixation: Tissues can be fixed in a wide range of fixatives. One of the more commonly used fixatives is an aldehyde (i.e.: glutaraldehyde) followed by osmium tetroxide. Dehydration: There are many different dehydration schedules that can be followed. A typical one is as follows: 70% Ethanol for 10 minutes 100% Ethanol for 10 minutes 100% Ethanol for 15 minutes 100% Propylene Oxide for 15 minutes 100% Propylene Oxide for 15 minutes **NOTE: Longer times may be required for some samples. Mixing Instructions: PROCURE-812 25ml Araldite 502 15ml DDSA 55ml DMP-30* 1.5-1.9ml *For better penetration and stability BDMA is recommended in place of the DMP-30. The quantity of BDMA which is required is 2.4-2.8ml. Slight variations of the accelerator (DMP-30 or BDMA) will drastically affect the colour and brittleness of the block. (FOR LARGER BATCHES INCREASE EACH COMPONENT PROPORTIONALLY) Prior to measuring and mixing, the resin and the anhydride should be warmed (60°C) to reduce their viscosity. Thorough mixing is imperative to be able to achieve uniform blocks. Although the mixture can be stored for up to 6 months at 4ºC it is highly recommended that freshly prepared embedding medium always be used. If you choose to store the mixture you should warm it thoroughly prior to adding the accelerator. Infiltration: It is recommended that for all of the infiltration steps a specimen rotator be used. Drain the tissue of most of the propylene oxide, leaving a little so the tissue does not dry out. Replace the solvent with a 1:1 solution of propylene oxide embedding medium and allow it to stand for at least 1 hour at room temperature. Remove the mixture, replace it with 100% embedding medium and leave for 6-12 hours at room temperature. Replace the old mixture with fresh embedding medium and allow it to sit for at least 2 hours. Embedding: This may be done in embedding capsules or a flat embedding mould. Transfer each sample to a dry capsule or mould and fill the mould with embedding medium. Cure the medium in an oven at 60ºC for 12 hours or until it is hard. Better sectioning properties of certain samples may be achieved if a time of 24 hours in the oven is used. Blocks can be trimmed and sectioned after the blocks return to room temperature. REFERENCES: Mollenhauer, H.H. (1964), Stain Technology, 39, 111.
Ruthenium Tetroxide 0.5% Aqueous Solution
EMS20700-05 In its crystal form, ruthenium (VIII) oxide (RuO4), is a golden yellow, volatile solid which sublimes at room temperature. It has a melting point of 25.4°C and a boiling point of 40°C. It is sparingly soluble in water (2% w/v at 20°C), but freely soluble in carbon tetrachloride. Solvents such as ether, alcohol, benzene and pyridine react violently with RuO4. Ruthenium tetroxide is not only less volatile and less toxic than osmium tetroxide but it is also a stronger oxidising agent. It reacts with many organic compounds like olefins, sulfides, primary and secondary alcohols, and aldehydes. It also degrades benzene rings. Applications and Instructions: Ruthenium tetroxide is closely related to osmium tetroxide and it is useful as a staining agent for electron microscopy of polymers and their blends, and as a fixative for biological samples. It fixes the membranes in rat kidneys, liver, and ventral lobe of prostrate, and these membranes appeared thicker than those preserved with other fixatives. Ruthenium tetroxide shows excellent staining of saturated and unsaturated polymer materials with improved image contrast. RuO4 staining also has a stabilising effect against electron beam damage of material films. Ruthenium tetroxide penetrates tissue very slowly, reacting strongly with proteins, glycogen, and monosaccharides. A typical procedure consists of prefixing with buffered 4% glutaraldehyde, followed by post fixation with buffered ruthenium tetroxide 0.1-0.5% at a pH=7.1 for 1 hour at 40°C. Specimens should be rinsed in water before dehydrating with alcohol or acetone. Note: Ruthenium tetroxide is a strong oxidizing agent and must be stored in the refrigerator away from direct sunlight. It reacts violently with filter paper and alcohol. Do not place ruthenium tetroxide solutions into waste containers containing alcohol, ether benzene, pyridine, or other organic compounds. Ruthenium tetroxide has an acrid odor. The vapors are irritating to eyes and the respiratory tract. Wear protective goggles and gloves and handle only in a fume hood. In case of spillage, flush with sodium bisulfite solution to decompose RuO4 and then flush with plenty of water.
Shield Mount with DABCO (IFMD) Mounting Medium
EMS17985-09, EMS17985-100 EMS Shield Mount with antifading agent 1, 4-Diazobicyclo-(2,2,2-octane (DABCO™) is an aqueous non-permanent, low viscosity mounting medium for immunofluorescence. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine and Redox. The fluorescence is retained during storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes. Applications Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium for Immunofluorescence. Instructions For Use Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium and apply coverslip carefully without getting air bubbles. The specimen is ready for visualisation under a microscope. The coverslip may move, so it may be necessary to seal the edges of cover slip with nail polish, any organic medium or our Limonene mounting medium. Store slide at 2-8°C in the dark. Handling And Storage 2-8°C is recommended. Protect from light, DO NOT FREEZE. Removal of Coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) distilled or deionised water for several minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
Shield Mount with DABCO™
EMS17985-150, EMS17985-200 EMS Shield Mount with antifading agent 1, 4-Diazobicyclo-(2,2,2-octane (DABCO™) is an aqueous non- permanent, low viscosity mounting medium for immunofluorescence. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine and Redox. The fluorescence is retained during storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes. This mounting medium is fortified with DAPI which is a counter-stain for DNA. This product is to be used in situ hybridisation techniques or other methods where fluorescence of DNA staining is required. DAPI excites at 360nm and emits at 460nm, producing a blue fluorescence. RNA is also stained with DAPI Applications Immunofluorescence, confocal microscopy. Reagent Ready to use mounting medium for Immunofluorescence. Instructions For Use Bring the vial to room temperature. Rinse slide to be mounted with DISTILLED OR DEIONISED WATER, touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium and apply coverslip carefully without getting air bubbles. The specimen is ready for visualiSation under a microscope. The coverslip may move, so it may be necessary to seal the edges of cover slip with nail polish, any organic medium or our Limonene mounting medium. Store slide at 2-8°C in the dark. Handling And Storage 2-8°C, Protect from light, DO NOT FREEZE. Removal of Coverslip Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) distilled or deionized water for several minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
Shield Mounting Medium with DAPI and DABCO™
EMS17989-20, EMS17989-21 Shield Mounting Medium with 4,6-diamidino-2-phenylindole (DAPI) and DABCO™ Fluoroshield with DAPI is an aqueous mounting medium for preserving fluorescence of tissue and cell smears. This unique formula prevents rapid photobleaching of FITC, Texas Red, AMCA, Cy2, Cy3, Cy5, Alexa fluoro 488, Alexa fluoro 594, Green fluorescent protein (GFP), tetramethyly rhodamine, Redox. Phycoerythrin (RP-E), Phycocyanin (PC), and Allophycocyanin (APC). Fluorescence is retained during prolonged storage at 4°C in the dark. This medium does not contain phenylenediamine, which destroys immunofluorescence of Cy dyes, RP-E, PC and APC. This mounting medium is fortified with DAPI which is a counter-stain for DNA. This product is to be used in situ hybridisation techniques or other methods where fluorescence of DNA staining is required. DAPI excites at 360nm and emits at 460nm, producing a blue fluorescence. RNA is also stained with DAPI. May encounter problems with frozen brain or other frozen tissues with lots of fat. Intended Use Immunofluorescence, confocal microscopy Reagent Ready to use mounting medium Refractive Index 1.364 ± 0.002 (This number applies to this mounting medium in solution. Refractive indexes change when the water solvent evaporates and mounting media dries on slides. We do not have the means to measure the refractive indexes of dry mounting mediums; however, we expect the numbers to go higher when dried. The refractive index of water is 1.3330.) Storage 2-8°C, Protect from light, DO NOT FREEZE Procedure Bring the vial to room temperature. Rinse slide to be mounted with distilled or deionised water; touch the edges of slide on a paper towel to remove excess water. Place slides on a flat surface away from light. Turn the vial upside down and open the dropper to remove any air bubbles. Apply 2-3 drops of mounting medium directly on top of the specimen. Let stand at room temperature for about 3-5 minutes in the dark. Apply cover slip, carefully avoiding air bubbles. The specimen is ready for visualisation under a microscope. One can seal the edges of the cover slip with nail polish or any organic mounting medium. If a coverslip is not sealed, air bubbles will appear in few days. Method for applying Coverslip: Put 1-2 drops of FLS on the specimen. After 3-4 minutes apply coverslip carefully avoiding air bubbles. Put Kimwipes on the top of coverslip, press gently to remove excess mounting medium. With 200 micropipette add organic mounting medium to seal the edges. Incubate at 37°C for one hour in the dark to dry organic mounting medium. For long term storage it is recommended that the slide be stored in the dark at 2-8°C. Removal of Coverslip: Coverslip can be removed before sealing the edges. Soak slide in warm (37°C) water for a few minutes. Carefully and slowly move the coverslip. Soak in water for an additional few minutes to remove coverslip. Rinse slide several times with warm water to remove all mounting medium. The slide can be remounted again.
Silica Gel Pellets
What is silica gel and how does it work? Silica gel is a solid desiccant (drying agent). It is not a wet gel as the name might suggest, and should not be confused with silicone gel. Silica gel is the most commonly used desiccant for providing efficient protection against excessive humidity in sealed spaces. This granular, vitreous, highly porous form of silica is made synthetically from sodium silicate. Silica gel beads contain millions of tiny pores that adsorb and hold water vapour and remain dry to the touch, even when fully saturated. This highly porous substance can adsorb up to 40 percent of its own weight at 50% RH and 25°C, and can take the relative humidity (RH) in a closed container down to about 40%, which makes this the highest water holding capacity, of any commercially used desiccant available. The adsorption up to 50% RH is a near‐linear relationship between the percentage of water uptake and the equilibrium relative humidity. This refined form of silica is produced in granular or beaded forms. Beaded silica gel has a higher mechanical strength than the granular form and is therefore less likely to generate dust. Also, the uniform bead size reduces gas flow channelling and minimises pressure drop. Silica gel is nearly harmless, which is why you find it in food products. Silica, or silicon dioxide (SiO2), is the same material found in quartz. Silica gel is essentially very porous sand. Some types of silica gel will "pop" when exposed to enough water. It can be reactivated and reused by heating. Silica gel is produced synthetically by the reaction of a mineral acid, usually sulphuric, and a sodium silicate solution to form a gelatinous precipitate that is washed, and then dehydrated to produce colourless silica gel. Ammonium tetrachlorocobaltate(II) (NH4)2CoCl4 or cobalt chloride CoCl2 may be added to silica gel as a moisture indicator. These indicators change colour depending on hydration. Cobalt Chloride is blue when dehydrated and pink when saturated. Orange silica gel desiccant is the latest alternative to blue indicating silica gel (cobalt chloride). Orange silica gel is impregnated with a safe organic indicator that displays an orange/yellow colour when active and changes to green when the desiccant becomes saturated. Silica gel traps water molecules inside its pores yet remains dry and appears physically unchanged. This synthetic form of sodium silicate and a mineral acid produces an amorphous, micro‐porous structure with a distribution of pore opening sizes of roughly 0.3‐6 nanometres. Comprised of tiny interconnecting pores, it absorbs water vapour by physical means, not a chemical reaction. Physical adsorption involves relatively weak intermolecular forces (van der Waals forces and electrostatic interactions) between the moisture and surface of the desiccant. Chemi Sorbents, such as calcium oxide, involve an actual chemical bond. Physical adsorption of moisture is typically exothermic. The strength of the adsorptive bonds can thus be measured by the heat of adsorption. The higher the heat of adsorption for moisture on the desiccant, the stronger the bonding and the less easily that moisture can be subsequently removed. These interconnected pores form a vast surface area (around 800 m²/g) that will attract and hold moisture inside the silica gel's crystalline structure by adsorption and capillary condensation, allowing silica gel to absorb up to 30‐40% of its weight of water. The adsorption capacity depends on a linear relationship between the percentage water uptake and the equilibrium relative humidity. Much of silica gel's popularity is due to its non‐corrosive, non‐toxic nature. Hence, non‐coloured silica gel has received US government approval for use in food and drug packaging. Beaded silica gel has a higher mechanical strength than the granular form and is therefore less likely to generate dust. However, silica gel is not considered biodegrade in water or soil. Silica Gel will continue to absorb moisture below freezing, but performs best at room temperatures. Advantages Visual indication of beads' saturation Environmentally friendly organic colour impregnation Frequently used in food and pharmaceutical applications High crush strength and high porosity Remains dry and free flowing, even when saturated Chemically inert, non‐corrosive and non‐toxic Odourless Does not form any by‐products Extremely large internal surface area Meets all Military Specifications for desiccants (Mil‐D‐3464, Type I & II) Available in any desiccant raw material (i.e. silica gel, clay, molecular sieve and others) Prevents mould, mildew, rust and corrosion Adsorbs moisture and odour inside a packaged environment Non‐Tearing, Non‐Dusting & Non‐Linting packets Compliant with FDA and USDA guidelines for packaged desiccants Available in Tyvek®, Non‐Woven, Kraft, Clear Poly & Sewn Cloth bags Silica gel applications Silica Gel has numerous uses, many of these in the rubric: the prevention of moisture damage. Desiccant Products, such as silica gel, activated clay or molecular sieves are drying agent used to combat humidity or moisture degradation. Products vulnerable to moisture damage need special protection during transportation, storage, etc. to ensure that it remains effective. Silica Gel packets and clay desiccant's provide a simple, dependable, and economical solution for preventing moisture damage in sealed packaging. Humidity in the space turns into condensation which can often cause irreparable damage. Our silica gel products will help to eliminate rust, corrosion, oxidation and tarnish on tools, jewellery, silverware, coin collections, musical instruments and much more. In addition, silica gel can protect seasonal storage, collectibles, leather, clothing and more from mildew, mould, fungus and odours. Furthermore, photographs, documents, and stamp collections can be protected from staining, spotting and cracking. Many manufacturers include silica gel packets in their electronics packaging which helps protect the electronics from moisture damage. Silica gel decreases moisture in the air which can limit the growth of mould and reduce spoilage. Silica gel may be used to dry electronic devices which had become wet. Placing the device in an airtight container with some silica gel, would remove most moisture, more effectively than air drying and more safely than heat. Storing electronic instruments in a sealed environment (plastic bag) with some silica gel prevents condensation and corrosion, which might cause serious damage. Vitamins and pills are often packaged with silica gel packets to prevent damage or decomposition due to moisture in the bottle. If a bottle of vitamins contained any moisture vapour and was cooled rapidly, the condensing moisture would ruin the pills. Silica gel packets are also used to help keep dried food, like beef jerky and pepperoni, fresh. In museum and library exhibitions and storage, silica gel is used as a preservation tool to control relative humidity (RH) invaluable air tight frames. Leather goods are often transported with silica gels to help protect them from moisture damage. Storing leather goods with silica gel packets can also help protect them from moisture and make them last much longer. If you have ever seen dried flowers and wondered how they managed to dry them and maintain the colours and shapes, they probably used silica gel. Covering the flowers with loose silica beads while they are still vibrant and firm dries them quickly, which helps maintain the colour and shape of the flowers. Silica gel can also be placed in a gun safe to help prevent moisture damage to both guns and ammunition. Dry bags or dry boxes used in water sports, should contain silica gel to dry bags or dry boxes and absorb any moisture. Adding a silica gel packet to an underwater camera case absorbs moisture from within the case and limit or eliminates condensation from the camera's window. Silica gel is most commonly encountered desiccant in everyday life, as clear beads packed in a semi‐permeable plastic. In this form, it is used as a desiccant to control local humidity to avoid spoilage or degradation of some goods. Because of poisonous doping agents (see below) and their very high adsorption of moisture, silica gel packets usually bear warnings for the user not to eat the contents. If consumed, pure silica gel is unlikely to cause acute or chronic illness, but if more than just a few beads are ingested the drying properties could cause some complications. Food‐grade desiccant should not include any poisons which would cause long‐term harm to humans if consumed in the quantities normally included with the items of food. Silica in this form can be easily purchased for applications such as keeping tools rust free in damp environments, long term storage, and preservation of dried food. Silica gel can be used anywhere, but it is really only effective in an enclosed environment. In a situation where new air (and moisture) is frequently added, an impractical large amount of silica gel and frequent changes to dried silica gel would be required to keep relative humidity at low levels. Applications: See the list below for other areas where the silica gel dehumidifiers can be helpful in eliminating moisture. Depending on application, different volume or mixed silica gel may be packaged inside the following substrates; Tyvek, Non‐Woven, Plastic Film, Paper, Sewn Cloth, or Reinforced Paper. Tool boxes ‐ help prevent rust in damp environments Storage containers Safes and vaults Silver drawer Firearms Photos Stored vehicles Cargo holds on RVs, boats, etc Collectibles (stamps, coins, etc.) Computers Metal and machine parts Industrial equipment Pharmaceuticals and vitamins Electronics Organic Liquid Drying Insulated Windows Flower Drying Telephone Cable Splicing Indication of Relative Humidity Insulated Glass Natural Gas Dehydration Military instruments Circuit Boards Documents Instruments Photographic equipment and film Foodstuffs Batteries Pet foods Optical devices Medical equipment Leather products Textiles Seeds Jewellery Packaged Gas & Liquid Dehydrators Hydrocarbon Recovery Units High Pressure Gas Drying Cartridges Cryogenic Purification Liquid Petroleum Gas Dryers Dynamic Drying of Refrigerants Does silica gel come ready to use? Yes. The silica gel does not need to be 'generated' before use. We sell it in a dry, airtight container, ready for use. What to do if silica gel has been ingested? We recommend contacting your local Poison Control Centre or seeking medical advice/attention. You can contact the Queensland Poisons Information Centre at 13 11 26 and they will provide you with advice based on who or what ingested the silica gel and how much of it. Based on this information, they will provide advice based on your situation. Silica gel is a non‐toxic material. See also http://www.health.qld.gov.au/PoisonsInformationCentre/ Is silica gel hazardous? Silica gel is inert, non‐toxic, non‐flammable, non‐reactive, and stable with ordinary usage. The substance was in existence as early as the 1640s as a scientific curiosity and is safe to use for protecting foods, medicines, sensitive materials and much more. It will react with hydrogen fluoride, fluorine, oxygen difluoride, chlorine trifluoride, strong acids, strong bases, and oxidisers. Silica gel is irritating to the respiratory tract, may cause irritation of the digestive tract, and dust from the beads may cause irritation to the skin and eyes ‐ so precautions should be taken. Some of the beads may be doped with a small amount of a moisture indicator, such as cobalt(II) chloride, which is toxic, and may be carcinogenic, and irritating to skin and the respiratory tract. Cobalt (II) chloride is deep blue when dry (anhydrous) and pink when moist (hydrated). We, along with several other retailers, offer a safer alternative which is an indicating silica gel that changes from orange to green and does not contain cobalt chloride. Note: Crystalline silica dust can cause silicosis but synthetic amorphous silica gel is non‐friable, and so does not cause silicosis. What is indicating silica gel? Indicating silica gel, was clear silica gel which had at least some of the beads impregnated with a chemical that changes colour with moisture content. When the colour has changed, the silica gel is at least partially saturated and needs to be replaced or reactivated by drying. This can be useful when long term humidity control is needed, like in a display case. What is "Saturation" and "Equilibrium Capacity"? Technically, for most practical purposes, these two terms cover the point at which a desiccant no longer absorbs moisture. Saturation is when the desiccant is full of water vapour and cannot adsorb more, even if there is excess moisture present. Very dry silica gel can reduce relative humidity further than silica gel which already carries some water. Equilibrium capacity applies to desiccant which has acquired so much moisture that the air retains an equally strong hold on water molecules as the desiccant. Adding more desiccants, of equilibrium capacity, cannot reduce relative humidity any lower. Exchanging the desiccant with very dry material is the obvious solution. What is Relative Humidity? Air always contains water molecules. At any given temperature the saturation point is where no more moisture is retained and excess moisture is condensed. The relative humidity of air is the moisture content expressed as a percentage of this saturation content. Silica gel is used to restrain relative humidity to below 50% of saturation capacity. At these low levels, mould growth and corrosion will not be promoted. Since the saturation point is temperature dependent, a drop in temperature may cause an increase in relative humidity. Disposal of silica gel products? The Silica gel can simply be thrown into the normal garbage. If the silica gel has come into contact with a toxic or hazardous material, it could potentially adsorb the material and become toxic. In this event, the silica gel should be disposed of in the same manner as the hazardous material that contaminated it. How much silica gel should be used in various applications? These are estimates and relate to situations when used in an airtight container. The amount of silica gel or desiccant required will depend on several factors including: volume contents of container, the chemical characteristics of the product, physical properties of the container, and conditions in which container will be stored and used. The amount will depend on the application however for a well sealed item, it is suggested that 200 grams of silica gel is needed for every cubic metre of volume of the package ‐ assuming that specimen and space were moderately dry when the silica gel was added. How does moisture cause damage to a product? Moisture trapped within a product package or leaking into it during storage and shipping can result in harm to the item. The maintenance of their physical product integrity, as well as the stability of many pharmaceutical formulations and diagnostic reagents, is often closely tied to the moisture conditions of the package environment. In poorly sterilised situations, moisture may promote the growth of moulds, mildews and fungi. Products that use polymers are prone to swelling in high humidity conditions, due to water causing intermolecular bonds between polymer chains to weaken. If a very water soluble solid, such as sugar coating, is exhibited to the right conditions, dissolution can trigger irreversible water uptake and subsequent deliquescence. Moisture in electronics causes corrosion and may lead to "short" circuits and other faults. Usually this will not produce sparks, the effects are more subtle, and the device may operate unreliably ‐ sometimes it work, sometimes it doesn't. A situation like this is frustrating, time‐wasting and may ruin an expensive instrument. Difference between silica gel and clay desiccants? Both bentonite clay and silica gel are desiccants, but there are major differences. Bentonite clay emits dust, whereas silica gel produces minute levels of dust particles. This addition renders silica gel the best choice to be packaged with items that could be adversely affected by dust particles like food or electronics. Silica gel is commonly packaged in cotton and perforated plastic. Silica gel adsorbs water up to 104°C, beyond that point it releases absorbed moisture. Bentonite clay adsorbs water only to 49°C at which point it begins to release any moisture back into the air. Both silica gel and bentonite clay can be reactivated for reuse. Bentonite clay and silica gel are used in industry, but silica gel is also commonly used in commerce and for domestic applications. Silica gel's absorbent capacity is 40% of its weight; bentonite clay's adsorbent capacity is only 25%. Silica gel's high adsorption capacity is the greatest available and considering its other desirable attributes (low dust, low toxicity and ability to absorb at higher temperatures) makes it the most popular desiccant. The buffering capacity of silica gel is determined by measuring the moisture gained or lost by silica gel as the RH changes in the surrounding air. For museum applications, where the case will cycle within a controlled RH range, the buffering capacity of silica gel is measured by its moisture holding (MH) value, which takes into account changes in buffering capacity based on whether the RH increases or decreases. Regular density silica gel is the most common type of silica gel and is universally available. RD silica gel is a form of silica manufactured from sodium silicate and sulphuric acid. Like clay, silica gel is non‐hazardous and is capable of adsorbing 40% of its weight in water vapour at 100% humidity. Because of its capacity for high moisture uptake in the low RH range, it is a very effective desiccant. For museum applications, the most important difference is buffering capacity, defined by the gel's MH value. Because of its poor hygroscopic properties around 45‐50% RH and above, it is not recommended for museum applications requiring control in the mid to upper RH range. Within the range of 40‐55% RH, it has an MH value of 2. Institutions located in areas with well defined wet and dry seasons, the buffering capacity of silica gel may become insufficient and the silica gel employed in sensitive cases may require replenishing. Recording a tray's silica gel net weight before and after drying will give a record of moisture levels. Silica gel has a porous molecular structure that closely resembles a sponge, and has the highest capacity of any commercial desiccant for moisture adsorption. Molecular Sieve is a manufactured crystalline version of Zeolite containing a network of uniform pores and empty cavities. Molecular sieve is derived from sodium, potassium or calcium alumina silicate. It is a non‐hazardous material. Molecular sieve is the desiccant of choice for the most demanding applications; so it is used to remove traces of moisture from absolute ethanol or acetone. Molecular sieve is the most aggressive and expensive of the primary desiccants. Reactivating/ drying of silica gel Silica gel products can be dried and reused "as new". Orange silica gel: Dry pellets are orange and turn dark green when near saturation. Blue indicating silica gel: Dry it's dark blue and with increased water uptake it becomes more pale and pink. The most efficient method to reactivate silica gel is with heat. When heat is applied to saturated silica gel, the beads will restore to the previous moisture absorbing capability and return to the original orange colour. Unlimited regeneration cycles are possible. Silica gel has a very high melting temperature, 1600°C. However, it will lose its chemically bound water and hygroscopic properties if heated above 300°C. In addition, there is a new class of indicator gels, incorporating organic dyes that are heat sensitive and their colour indicating dye will be affected above 125‐150°C . Therefore, it is not recommended that orange indicating silica gel is heated above 120°C. Also, if heated above 120°C through several reactivation cycles, the material may disintegrate into powder. This in turn, reduces the adsorption capacity of the material and may eventually lead to the loss of indicating colour within the crystals. The minimum heat necessary should be used when removing moisture from silica gel; ProSciTech recommends regeneration at between 105 °C and 120 °C. This will prevent the silica gel from deteriorating and it may be re‐generated numerous times. Blue indicating silica gel has a higher tolerance and may be dried by heating it to 150°C. Lower heat for regeneration requires longer drying times, but the degradation of silica gel is reduced. Increasing heat vapourises adsorbed moisture and above 105°C is removes all water molecule from silica gel. A porous desiccant like silica gel, removes water from the surrounding air by two mechanisms: multi‐layer adsorption and capillary condensation. Multi‐layer adsorption is the attraction of thin layers of water molecules to the surface of the desiccant. Since the desiccant is very porous, the surface area is high and significant amounts of water can be attracted and absorbed. Capillary condensation is when the smaller pores become filled with water. Capillary condensation occurs when saturated water vapour pressure in a small pore is reduced by the effect of surface tension. In a conventional oven, the time of regeneration varies from minutes to hours, depending on temperature and the thickness of the gel within a dish. Although silica gel can be dried in a microwave oven, it is difficult to determine the temperature inside the gel. Particularly the orange gel should not be dried using microwaves since excessive heating denatures the indicator dye. As metal cannot be used in a microwave oven, only glass, ceramic or microwave safe plastic with a high melting temperature should be used to hold the gel. Place 'used' silica gel beads into a large tray Use an oven at 105 ‐ 120°C for two hours, or continue heating for 30 minutes after the gel turned orange. Check the silica gels beads periodically. Caution:the gel beads are very hot to touch. Definitions: Absorption is when a substance is chemically integrated into another. For example, when acid is added to water, the acid is diluted and the water cannot be driven off by just heating. The acid cannot be concentrated by driving the water off. We can absorb knowledge as this becomes part of us. Adsorption applies when one substance is being held inside another by physical bonds only. The term is used in surface chemistry and physics. So, a film of moisture is adsorbed onto cold surfaces. The surface is not chemically changed and when the surface is heated, the adsorbed water will mostly volatilise. Most desiccants do not absorb water or other substances. Instead, they capture molecules by adsorption and capillary action, they sequester them. Use and applications of silica gel How long for silica gel will protect an enclosed area before it needs to be recharged depends on many factors. Deciding factors are the volume of the area, the amount of new, additional air penetrating the area, and if the contents was reasonably desiccated. If the area is not sealed tightly, outside air will infiltrate and shorten the time before the unit needs to be reactivated. Also, if you frequently break the seal and expose the area to new air, it will shorten the time until reactivation is needed. When silica gel is first placed into an enclosed area, it may be saturated quickly as it adsorbs residual moisture adsorbed on the specimens and chamber surfaces. After the residual dampness is removed, a dry condition can be maintained with less frequent need to reactivate the silica gel. The more air tight the environment is, the less reactivation times needed. Making a chamber airtight will limit the amount of moisture leaking into the area. Also, avoid frequent opening of the desiccated space. For example, silica gel will work well within a storage trunk, but it will not work as well if the lid is opened and closed several times a day. The adsorption capacity of silica gel can be destroyed by contamination with dust, grease or petroleum products, and also by heating the material above 125°C. How does silica gel control relative humidity? All silica gels are hygroscopic, responding to the relative humidity (RH) of the surrounding in the same way as most organic materials, such as paper and wood. The amount of moisture in silica gel will increase as the RH rises, and will decrease when the RH falls. Unlike organic materials that expand and contract with changes in moisture content, silica gel volume remains unchanged. In a near‐airtight space like a display case, a quantity of silica gel acts as a buffer hydrating and dehydrating the case's RH. Additionally, silica gel adsorbs and desorbs much larger amounts of moisture when the RH changes under normal conditions. Storing silica gel Most damage to stored supplies is caused by humidity trapped within the enclosed storage unit itself. The crucial factor is to keep it in a sealed, air‐tight environment until it is needed for use ‐ it will, of course, adsorb moisture from any environment. Although the moisture uptake rate is not fast, silica gel can be particularly vulnerable to poor storage conditions. It is recommended that they are not left open to the atmosphere for longer than 15 minutes. Useful life of silica gel http://www.howstuffworks.com/question206.htm http://www.theruststore.com/Silica‐Gel‐FAQ‐W40C2.aspx http://www.apsnyc.com/html/humidity_control_FAQ.htmL http://en.wikipedia.org/wiki/Silica_gel http://www.brownell.co.uk/products/category/47 http://www.geejaychemicals.co.uk/faq.htm http://www.apsnyc.com/pdf/silica_gel_SW_2003.pdf http://www.replen.com.sg/our‐products/dessicants/orange‐environmental‐silica‐gel/ http://www.sorbentsystems.com/desiccants_terminology.htmL http://www.silicagelpackets.com/faqs/ http://www.desiccare.com/faq.htm http://talasonline.com/photos/instructions/silica_gel_info.pdf http://talasonline.com/photos/instructions/silica_gel_FAQ.pdf http://talasonline.com/photos/instructions/silica_gel_reconditioning.pdf http://www.chemspider.com/Molecular‐Formula/C25H30ClN3
Silver Conductive Coating
EMS12684-15 – Silver conductive 18DB70X coating (DG) EMS Silver Conductive Coating is a highly conductive acrylic paint designed to take conductive paths or reduce electromagnetic or radio frequency interference (EMI/RFI). Because it is a durable acrylic resin, long-term protection from EMI/RFI is assured. It minimises loss of metallisation through rubbing, and by the oxidation resistant silver that slows down conductivity degradation with age. The flake shape helps ensure maximum points of contact to ensure better conductivity. Loss of shielding through paint peeling is unlikely since the acrylic resin system has been UL-tested and shown to adhere to even difficult substrates like AMS and polycarbonates. The primary application is to provide an excellent conductivity EMI/RFI shielding suitable for harsh environments. It may also act as a conductive base for applications where it is necessary to impart the highest degree of conductivity to a surface. Being non-magnetic, it also offers a low relative permeability that provides reasonable skin depths, which makes it ideal for microwave transmissions applications. Feature Highlights and Benefits Environmentally meets RoHS directive for Low-VOC Stronger adhesion than water-based coatings Rub-off resistant Repairable and removable thermoplastic paint system Tough and durable coat with excellent weatherability Corrosion resistant coating: Salt-Spray Tested Meets MIL-STD-883H (Volume Resistivity = 0.0002Ω·cm) High Surface Conductivity (≥15 Siemen)–Low Surface resistance of 0.066Ω /sq @ 1 mil Median attenuation 75 dB ±20 dB per 25.4 µm (~1.0 mil) for frequency range of 10 MHz to 18 GHz Cure Rates and Shelf Life *Dry to Touch (Liquid): 3 to 5 min *Recoat time (Liquid): 2 min Full Cure at room temp.: 24 hour Full Cure at 65°C: 1 year **Storage Temperature Limits: -5 to +40°C *Assumes let 1.00:0.75 let down with thinner. **The product must stay within storage temperature limits stated. Service Ranges Service Temperature: -40 to +120°C ***Maximum coverage per 900ml: <168 000 cm² ***Maximum coverage per US gal: <709 000 cm² ***Idealised estimate based on a coat thickness of 25 µm (1.0mil) and 65% transfer efficiency. Principal Components and their CAS Number Silver: 7440-22-4 Acrylic Resin: 9003-01-4 Acetone: 67-64-1 Ethanol: 64-17-5 Toluene: 108-88-3 Properties of Cured EMS Conductive Silver Coating NOTE: The first coat thickness is typically around 25 µm [1.0 mil]. Electric & Magnetic Properties Method Value Volume Resistivity (Tested by an external and independent laboratory using four point probe.) Method 5011.5 in MIL-STD-883H 0.0002 Ω·cm Surface Resistance :1 x coat @ 1 mil square probe *Resistance *Conductance 0.066 Ω/sq 15S :2 x coats @ 2 mil " 0.055 Ω/sq 18S :3 x coats @ 2.5 mil " 0.040 Ω/sq 25S *NOTE: Surface resistance is given in /sq and the corresponding conductance in Siemens (S or ) Magnetic class relative permeability Diamagnetic (Non-magnetic) <1.0 Shielding Attenuation 33 µm [1.0 mil] 10 to 100 kHz IEEE STD 299-1997 54 dV to 75 dB 100 kHz to 1 MHz " 50 dB to 65 dB 1 MHz to 10 MHz " 54 dB to 65 dB 10 MHz to 100 MHz " 41 dB to 54 dB 100 MHz to 1 GHz " 35 dB to 67 dB 1 GHz to 10 GHz " 41 dB to 59 dB 10 GHz to 18 GHz " 36 dB to 72 dB Physical Properties Method Value Resin technology – Lacquer (Thermoplastic) Colour Visual Metallic Silver Grey Abrasion resistant – Yes Blister resistant – Yes Peeling resistant – Yes Environmental & Aging Study Method Value Salt Spray Test: 7 day @35 °C +Salt/Fog ASTM B117-2011 Cross-hatch adhesion ASTM D3359-2009 5B = 0% area removed Cracking, unwashed area ASTM D661-93 None Visual Colour, unwashed area ASTM D1729-96 Severe yellowing & discoloration Peeling, unwashed area ASTM D1729-96 None Graph–Silver Coating Surface Resistance at Different Thicknesses Properties of Uncured EMS Conductive Silver Coating Physical Property: Mixture Colour: Metallic silver grey Density @ 25°C2.15 g/mL Solids Percentage (wt/wt) (Percentage for liquid only before thinning)~73% Viscosity at 25°C (Brookfield viscometer)~8,000 cP Flash Point-16°C Odour Ethereal, benzene-like Compatibility Chemical The silver filler is very resistant to oxidation, except in environments containing contaminants such as ozone or H2S, which tarnish its surface. Silver oxide remains conductive so degradation due to oxidation is not as bad as it is with many other metal oxides. Common paint solvents like acetone, toluene, MEK, and xylene can dissolve the thermoplastic resin. This does make for easier coating repair and work characteristics but is also makes the coating unsuitable for environments that have a lot of solvents. Adhesion EMS Conductive Silver Paint coating adheres to acrylics, metals, epoxies, wood, ABS, PBT, PC, PU, and PVA. NOTE: It is not compatible with contaminants like water, oil, and greasy flux residues which may affect adhesion. If contamination is present, clean the surface before coating. Storage Store in a dry area, between -5°C and 40°C (23°C and 104°F).
Slide Coating Adhesive
Introduction: Prepared from 3-Aminopropyltriethoxysilane in Acetone. The adhesive is ideal for coating glass slides prior to adhering either paraffin or plastic sections to the slides. This adhesive prevents sections from falling off of the slide during processing. Instructions for use: Dip the slide in the Slide Coating Adhesive, using either our 5 or 10-slide unit Coplin Jar or the Dip Miser. Rinse the slide in 100% Acetone. Rinse the slide in distilled water. Dry the slide at 40°C in a dust-free desiccator. The slide is now ready to use. Tissue sections (0.25-0.5 microns) are transferred to a drop of water on the slide (coated). Place the slide on a hot plate at 70-80°C for 20-30 seconds or until the sections have expanded and they become flat. Dry the sections at 68-70°C for an additional 1-3 minutes. Then follow your protocol for etching and/or staining of the sections. CAUTION: Use I030 (Slide Coating Adhesive) under a fume hood for the fumes are toxic. Reference: David L. Mason et. al., The J. of Histotechnology/Vol. 15, No 4/1992.
Sodium Cacodylate Trihydrate
C020 & C0205 Sodium cacodylate trihydrate is the most favoured buffering vehicle used with fixatives in specimen preparation for electron microscopy. Also called cacodylate or cacodylic acid, this chemical avoids the microprecipitation on thin sections that can occur with phosphate buffers if the specimen is not well rinsed between pre- and post-fixation. The chemical does not form a precipitate in sea water. Ultrastructural preservation is excellent. After some years the chemical breaks down at room temperature, so storage in the refrigerator is recommended. Made up solutions should also be stored refrigerated where they last for many months before some growths occur. Note: This compound contains arsenic. Arsenic is poisonous and carcinogenic. It is absorbed through the skin. Avoid contact with the powder and solutions. Use gloves and fume-hood. Wash hands after use. Electron microscopists use only small quantities of this chemical. Clean working habits and simple precautions can make its use safe. Properties: C2H6AsNaO2.3H2O F.W. 214.02 CAS #124-65-2 Certificate of Analysis: Identity Conforming Appearance Small Crystals Colour White Odor None Aqueous solution (2g+10 ml) Colourless Assay 99.52% Loss on drying 25.39% Chlorides 0.0050% Sulphates 0.0150% Inorganic Arsenic 0.0030% For animal tissue, buffers are normally used at 0.1M and for plant tissue at 0.01 molar. To prepare the latter, the same stock solution of 0.2M may be used and is diluted 1:9 parts of water before use. To make up a 0.2 molar stock solution: Into a 100ml volumetric flask, with a magnetic stirring bar, weigh 4.3g of sodium cacodylate. Add double distilled water to 100ml mark. Stir to dissolve. Pour solution and stirring bar into a suitable container and adjust the pH to the desired reading (plant tissue most commonly pH 6.8, animal tissue pH 7.2) using 0.2N HCl. The natural pH of the solution is about 7.8. Most commonly the solution is mixed 1:1 to make: With water to make single strength buffer for storage and rinsing. With 5% glutaraldehyde to make 2.5% glutaraldehyde in single strength buffer. With 2% osmium tetroxide to make 1% OsO4 in single strength buffer.
Sodium Chloride (NaCl) Quality Specification
Product Specification Product name Sodium Chloride LR Alternate name(s) Table salt; sea salt; halite; rock salt Description Colourless transparent crystals or white crystalline powder. Hygroscopic. Properties Chemical formula NaCl Molecular weight 58.44 Product code C017 CAS No. 7647‐14‐5 General Information Soluble in glycerol. Slightly soluble in alcohol. Incompatible with strong acids and oxidising agents. Non combustible material; emits toxic fumes under fire conditions. Hazard and Safety Data UN group none allocated Class none allocated UN number none allocated Hazchem code none allocated CS MSDS code 1CH69 Poison schedule not scheduled Not classified as hazardous according to the criteria of ASCC. Quality Specification Assay 99.0% min. Specific Properties and Impurities [Typical levels]: Melting point 801°C Boiling point 1461°C (1013 hPa) Vapour pressure 1.3 hPa (865°C) Specific gravity 2.165 Solubility in water 358 g/L (20°C) Sulphate <0.2% Calcium <0.01% Iron <0.001% Arsenic <0.00005% Insoluble matter <0.03% Magnesium <0.01% Copper <0.0002% Cadmium <0.00002% Lead <0.0001% Mercury <0.000005%
Spurr's Embedding Kit
Introduction: Spurr's Low Viscosity embedding mixture is recommended because of its excellent penetration qualities, which provide good and rapid infiltration of tissues. It is easy to prepare, and mixes rapidly by shaking and swirling. The hardness is adjusted by changing the amount of the flexibiliser, DER 736 or DER 732; the blocks have good trimming and sectioning qualities, and the sections are tough under the electron beam. (Grids, without supporting membranes can be used.) Ingredients: ERL 4221 – The direct replacement for ERL 4206. DER 732 / DER 736 - (DER 736 can be harder to get in Australia), DER 732 has a slightly greater epoxy equivalent and requires 10% less accelerator in Spurr's formula. It is also is a flexible, low viscosity, light colour, epoxy resin for use in coatings and adhesives. This resin, with conventional bisphenol A based epoxy resins, imparts flexibility, elongation and improves impact resistance. NSA - Nonenyl succinic anhydride, a hardener with a relatively low viscosity of 102.8cP at 25°C, and a M.W. of 227. A minimum exposure to air is recommended to avoid hydrolysis. DMAE - Dimethylaminoethanol (S-1) an accelerator, used because of its low viscosity and results in blocks with less colour. In addition, it induces rapid cure when the temperature is elevated to 70°C. It is effective in a very low concentration. (less than 1.0%) The optimum concentration for colour transparency is 0.7-0.75%. Original Formulation using DER 736: See end of Document for special formulations ERL 4221 8.6ml DER 736 7.0ml NSA 24.2ml DMAE 0.34ml Cure Time at 70°C 8 hours Pot Life 3-4 days Amended Formulation using DER 732: See end of Document for special formulations ERL 4221 8.6ml DER 732 7.6ml NSA 24.2ml DMAE 0.30-0.31ml Cure Time at 70°C 8 hours Pot Life 3-4 days Mixing Instructions: Add each component in turn to a disposable plastic beaker. An exact weight is recommended, and care must be used in dispensing the final amounts of each component so that no excess is added. The catalyst (DMAE) should be added last, after gently mixing the three other components. The complete formula should be mixed thoroughly. The complete mixture with the hardener can be used immediately for infiltration, and then for embedding. Although the mixture can be stored in a disposable syringe, well capped and with no air, in a freezer for several months it is highly recommended that freshly prepared embedding medium always be used. If you choose to store the mixture, it is imperative that you allow it to come to room temperature prior to use to avoid atmospheric water condensation on the resin, which prevent proper polymerisation. Dehydration - Infiltration and Polymerisation: This embedding media is compatible with all dehydrating agents: acetone, dioxane, ethanol, hexyleneglycol, isopropyl alcohol, propylene oxide, tert-butyl alcohol. The schedule and concentration can be established by the investigator however for a rule of thumb we recommend the following: Dehydration is generally done at room temperature in a graded series of Ethanol starting at 50% then going to 70, 95 and 100% for no less than 20-30 minutes each step. All dehydrating agent must totally be removed during infiltration due to the fact that it will effect curing. The embedding media is completely compatible with ethanol. Thus, it is not mandatory to have a change to propylene oxide prior to infiltration as is true for other epoxy resin mixtures. If working with plant cells it is recommended to use propylene oxide. The infiltration (one should employ a specimen rotator) can be started by adding the embedding media to the dehydrating fluid left in the vial with the tissue (1:2). Swirl the mixture and allow it to stand for 2- 3 hours. Then replace with a 1:1 dehydrating agent/embedding medium, swirl and allow to sit overnight. Replace with a 1:3 dehydrating agent/embedding medium, swirl, and allow it to stand for another 2 to 3 hours. Pour and drain the mixture and add fresh embedding media (100% Spurr). For small specimens, 5-6 hours; for large specimens, 5-6 hours followed by overnight. Curing takes 16-24 hours at 60°C. (The mixture can be left in an oven overnight). Special Formulations: For Biopsies from Kidney, Liver, GI, Esophagus Mixing Directions : ERL 4221 18ml DER 736 14ml NSA 48ml DMAE 0.6ml Dehydration: 50%, 70%, 95% ETOH for 20-30 minutes each change 2 x 100% ETOH for 30 minutes Infiltration*: 1 part Spurr to 2 parts ETOH for 3 hours 1 TO 1 overnight/over weekend 3 to 1 (Spurr to ETOH) 3 hours 100% Spurr 6-8 hours Polymerisation: Then Polymerise overnight (12 hour Max) at 70-80°C *All infiltration Procedures shall be run on a rotator. For Biopsies from Muscle, Nerve or Skin Dehydration: 50%, 70%, 95% ETOH for 20-30 minutes each change 2 x 100% ETOH for 30 minutes Infiltration*: (Spurr/ETOH 100%) 1 part Spurr to 2 parts ETOH 24 hours (over weekend) 1 TO 1 24 hours (over weekend) 3 to 1 (Spurr to ETOH) 24 Hours 100% Spurr 24 Hours Polymerisation: Then Polymerise overnight (12 hour Max) at 70-80°C *All infiltration Procedures shall be run on a Rotator.
Spurr's Embedding Kit (Low Viscosity Embedding Media)
Spurr's Low Viscosity embedding mixture is recommended because of its excellent penetration qualities, which provide good and rapid infiltration of tissues. It is easy to prepare and mixes rapidly by shaking and swirling. The hardness is adjusted by changing the amount of the flexibilizer, DER 732; the blocks have good trimming and sectioning qualities and the sections are tough under the electron beam. (Grids, without supporting membranes can be used.) INGREDIENTS: ERL-4221 is a Cycloaliphatic Epoxide Resin. Specific Gravity (H2O=1): 1.173 @ 20°C/20°C Flash Point – Closed Cup: 118°C Boiling Point (760mm/Hg):>250°C DER 732 - diglycidyl ether of polypropylene glycol, a flexibilizer to control the hardness of the polymerized block. It was selected because of its low viscosity: 30-60 cP at 25°C. It has a M.W of 380 and an epoxy equivalent of 175-205. There is little difference between DER 732 and DER 736. The latter is used elsewhere but is not available in Australia NSA - nonenyl succinic anhydride, a hardener with a relatively low viscosity of 102.8cP at 25°C, and a M.W. of 227. A minimum exposure to air is recommended to avoid hydrolysis. DMAE - dimethylaminoethanol (S-1), an accelerator, used because of its low viscosity and results in blocks with less colour. In addition, it induces rapid cure when the temperature is elevated to 70°C. It is effective in a very low concentration. (less than 1.0%) The optimum concentration for colour transparency is 0.7-0.75%. ABCDMODIFICATIONS FIRM STANDARDHARDSOFTLONGER POT LIFE LOWER VISCOSITYERL 420610.0g10.0g10.0g10.0gDER 7326.0g4.0g8.0g6.0gNSA26.0g26.0g26.0g26.0gDMAE0.3g0.3g0.3g0.2gCURE TIME AT 70°C (hours)88816POT LIFE (DAYS)3-43-43-47 MIXING INSTRUCTIONS: Add each component in turn to a disposable plastic beaker. An exact weight is recommended, and care must be used in dispensing the final amounts of each component so that no excess is added. The catalyst (DMAE) should be added last, after gently mixing the three other components. The complete formula should be mixed thoroughly. The complete mixture with the hardener can be used immediately for infiltration, and then for embedding. Although the mixture can be stored in a disposable syringe, well capped and with no air, in a freezer for several months, it is highly recommended that freshly prepared embedding medium always be used. If you choose to store the mixture, it is imperative that you warm it thoroughly prior to use. DEHYDRATION - INFILTRATION AND POLYMERIZATION: This embedding media is compatible with all dehydrating agents: acetone, dioxane, ethanol, hexyleneglycol, isopropyl alcohol, propylene oxide, tert-butyl alcohol. The schedule and concentration can be established by the investigator. Dehydration is generally done at room temperature. All dehydrating agent must totally be removed during infiltration due to the fact that it will affect curing. The embedding media is completely compatible with ethanol. Thus, it is not mandatory to have a change to propylene oxide prior to infiltration, as is true for other epoxy resin mixtures. If working with plant cells it is recommended to use propylene oxide. The infiltration (one should employ a specimen rotator) can be started by adding the embedding media to an equal quantity (1:1) of the dehydrating fluid left in the vial with the tissue. Swirl the mixture and allow it to stand for 30 minutes to 2 hours. Replace with a 1:3 dehydrating agent/embedding medium, swirl, and allow it to stand for another 30 minutes to 2 hours. Pour and drain the mixture and add fresh embedding media. For small specimens leave 4-6hours; for large specimens leave 4-6 hours followed by overnight. Curing takes 16-24 hours at 60°C. The mixture can be left in an oven overnight. LOW VISCOSITY EMBEDDING MEDIA KIT CONSISTS OF: C056, ERL 4206 - vinyl cyclohexene dioxide - 225ml C047, DER 732 - diglycidyl ether polypropylene glycol - 200ml C059, NSA - nonenyl succinic anhydride - 500ml C050, DMAE - dimethylaminoethanol - 25ml REFERENCES: Spurr, A.R.(1969), J. Ultrastruct. Res. 26,31.
Sub-X® Mounting Medium
Sub-X Mounting Medium is a toluene-based low viscosity quick-drying permanent mountant that has been designed to be suitable with all xylene alternatives on the market. Can be used for any coverslip preparation that has been cleared in xylene or toluene. It is suitable for both manual and automated coverslipping. Sub-X contains an antioxidant that prevents stain fading. DO NOT USE Sub-X® clearant to dilute mounting medium. A small amount of Toluene or Xylene must be used to reduce viscosity. Xylene or toluene must be used to remove coverslips that have set up. Store at room temperature 15-30°C Flash point CC: 12°C Hazardous Ingredient % wt. CAS no. Toluene <75 108-88-3 Acrylic Resin <50 80-62-6 Butyl Benzyl Phthalate <10 85-68-7 Antioxidant <1 128-37-0
Sylgard™ 184 Silicone Elastomer
SYLGARD™ 184 Silicone Elastomer - Transparent encapsulant with good flame resistance Features and Benefits: Flowable Room temperature and heat cure Good dielectric properties Rapid, versatile cure processing controlled by temperature High transparency allows easy inspection of components Composition: Two-part 10 to 1 mix ratio Polydimethylsiloxane elastomer Applications: LED lighting encapsulation Power supplies Connectors Sensors Industrial controls Transformers Amplifiers High voltage resistor packs Relays Adhesive/encapsulant for solar cells Adhesive handling beam lead integrated circuits during processing Typical Properties Property Result One or two part Two Colour Colourless Viscosity (Base) 5100cP 5.1Pa-sec Viscosity (Mixed) 3500cP 3.5Pa-sec Thermal Conductivity 0.15btu/hr ft °F 0.27W/m °K Specific Gravity (Cured) 1.03 Working Time at 25°C (Pot Life - Hours) 1.5hrs Cure Time at 25°C 48hrs Heat Cure Time at 100°C minutes 35 35mins Heat Cure Time at 125°C minutes 20 20mins Heat Cure Time at 150°C minutes 10 10mins Durometer Shore A 43 Dielectric Strength 500V/mil 19kV/mm Volume Resistivity 2.9E+14Ωcm Dissipation Factor at 100 Hz 0.00257 Dissipation Factor at 100 kHz 0.00133 Dielectric Constant at 100 Hz 2.72 Dielectric Constant at 100 kHz 2.68 Linear CTE (by DMA) 340ppm/°C Tensile Strength PSI 980PSI 6.7MPa 69kg/cm2 Refractive Index @ 589nm 1.4118 Refractive Index @ 632.8nm 1.4225 Refractive Index @1321nm 1.4028 Refractive Index @ 1554nm 1.3997 Description: Dow silicone 10 to 1 encapsulants are supplied as two-part liquid component kits. When liquid components are thoroughly mixed, the mixture cures to a flexible elastomer, which is well suited for the protection of electrical/PCB system assembly applications. Dow silicone encapsulants cure without exotherm at a constant rate regardless of sectional thickness or degree of confinement. Dow silicone elastomers require no post cure and can be placed in service immediately following the completion of the cure schedule. Standard silicone encapsulants require a surface treatment with a primer in addition to good cleaning for adhesion while primerless silicone encapsulants require only good cleaning. HOW TO USE Application methods: Automated metered mixing and dispensing Manual mixing Substrate Preparation: Clean and degrease all surfaces with a suitable solvent prior to potting. Note: Make sure that all solvent is removed. Mixing: Sylgard 184 is packaged in lot matched kits with the base and curing agent in separate containers. Mix the two components thoroughly using a weight or volume ratio of 10:1. It is recommended to vacuum de-air, with a residual pressure of 10-20 mm mercury, which, when applied for applied for 30 minutes, will sufficiently de-air the material. How to Apply: Apply the encapsulant without entrapping any air. Note: Vacuum encapsulation is recommended for complex geometries. For information on appropriate dispensing equipment for your application, please contact Customer Service at 1-800-523-5874. Curing: Cure reaction begins with the mixing process. Initially, cure is evidenced by a gradual increase in viscosity, followed by gelation and conversion to a solid elastomer. Pot life is defined as the time required for viscosity to double after base and curing agent are mixed and is highly temperature and application dependent. Please refer to the data table. Cure using one of the following recommended schedules: 15 minutes at 150°C 1 hour at 100°C 4 hours at 65°C 24 hours at 23°C Note: Large components and assemblies may require longer curing times. Note: At 23°C the material will have cured sufficiently in 24 hours to be handled; however it takes 7 days for full mechanical and electrical properties to be attained. Temperature Ranges: For most uses, silicone elastomers should be operational over a temperature range of -45 to 200°C for long periods of time. However, at both the low and high temperature ends of the spectrum, behaviour of the materials and performance in particular applications can become more complex and require additional considerations and should be adequately tested for the particular end-use environment. For low-temperature performance, thermal cycling to conditions such as -55°C (-67°F) may be possible, but performance should be verified for your parts or assemblies. Factors that may influence performance are configuration and stress sensitivity of components, cooling rates and hold times, and prior temperature history. At the high-temperature end, the durability of the cured silicone elastomer is time and temperature dependent. As expected, the higher the temperature, the shorter the time the material will remain useable. Compatibility: In some cases, Sylgard 184 may not cure to optimum properties when in contact with certain plastics or rubbers. Note: Baking the substrate slightly above the cure temperature or cleaning it with solvent will normally eliminate the problem. Certain chemicals, curing agents and plasticisers can inhibit a complete cure, including sulfur, polysulfides, polysulfones and other sulfur containing materials; organo-tin compounds; silicone rubber containing organo-tin catalysts; and amines, urethanes, amides and azides. Repairability: In the manufacture of electrical devices and PCB system assemblies it is often desirable to salvage or reclaim damaged or defective units. With most non-silicone rigid potting/encapsulating materials, removal or entry is difficult or impossible without causing excessive damage to internal circuitry. Dow silicone encapsulants can be selectively removed with relative ease, depending on the chosen remove method and technique and repairs or changes accomplished, and the repaired area repotted in place with additional product. To remove silicone elastomers, simply cut with a sharp blade or knife and tear and remove unwanted material from the area to be repaired. Sections of the adhered elastomer are best removed from substrates and circuitry by mechanical action such as scraping or rubbing and can be assisted by applying DOWSIL™ OS fluids to swell the elastomer. Before applying additional encapsulant to a repaired device, roughen the exposed surfaces of the cured encapsulant with an abrasive paper and rinse with a suitable solvent and dry. This will enhance adhesion and permit the repaired material to become an integral matrix with the existing encapsulant. Silicone prime coats are not recommended for adhering products to themselves. Handling Precautions: PRODUCT SAFETY INFORMATION REQUIRED FOR SAFE USE IS NOT INCLUDED IN THIS DOCUMENT. BEFORE HANDLING, READ PRODUCT AND SAFETY DATA SHEETS AND CONTAINER LABELS FOR SAFE USE, PHYSICAL AND HEALTH HAZARD INFORMATION. THE SAFETY DATA SHEET IS AVAILABLE ON THE DOW WEBSITE AT DOW.COM, OR FROM YOUR DOW SALES APPLICATION ENGINEER, OR DISTRIBUTOR, OR BY CALLING DOW CUSTOMER SERVICE. Usable Life and Storage: Shelf life is indicated by the “Use Before” date found on the product label. Refer to the product label for storage temperature requirements. Special precautions must be taken to prevent moisture from contacting these materials. Containers should be kept tightly closed and head or air space minimised. Partially filled containers should be purged with dry air or other gases, such as nitrogen. Disposal Considerations: Dispose in accordance with all local, state (provincial) and federal regulations. Empty containers may contain hazardous residues. This material and its container must be disposed in a safe and legal manner. It is the user’s responsibility to verify that treatment and disposal procedures comply with local, state (provincial) and federal regulations. Contact your Dow Technical Representative for more information. Note: This product is neither tested nor represented as suitable for medical or pharmaceutical uses.
Tannic Acid (LMGG)
Introduction: Tannic acid was introduced as a secondary fixative mixture with aldehydes for biological tissues, and also as a stain. Specimens treated with Tannic acid show increased contrast and more delineation of cell membranes. A low molecular weight, Tannic acid (LMGG), galloylglucose (C14H1009)n, provides and overcomes the previous problems of unsatisfactory penetration, extraction, and precipitation when high molecular weight Tannic acid (C76H52O46) was utilised. It works primarily as a mordant between osmicated structures and Lead citrate of the post-staining, revealing additional ultra-cellular structures and details better delineated. Procedure: The procedure reported by Simionescu (1976) involves the following steps: (Sodium Cacodylate buffer is preferred.): Fix tissue in Glutaraldehyde and Osmium tetroxide. Rinse in 0.1M buffer (pH 7.2) 3 times for five minutes each time at room temperature. Treat with 1% LMGG in 0.5M buffer for 30 minutes. Rinse in the same buffer containing 1% Sodium sulphate for 5 - 10 minutes. Dehydrate in ethanol followed by Propylene oxide: leave over night in a 1:1 EMbed 812 and Propylene Oxide mixture at room temperature; embed the next day. Stain thin sections in Lead citrate for 3 - 5 minutes. Solution: 1% LMGG Tannic acid (C14H1009) in 0.005M Sodium Cacodylate buffer, freshly prepared. Concentration of LMGG can be adjusted in a range of 0.25% to 2.0% according to the nature of the tissue and section thickness. Reference: N. Simionescu and M. Simionescu, J. Cell Biology (1976-70), 608-621.
Technovit 9100 MMA Embedding Kit
Components Technovit 9100 Basic Solution - Component 1 SDS The Technovit 9100 basic solution is comprised of stabilised methyl methacrylate. The hydrophily is improved through the addition of a suitable hydrolysing agent. Technovit 9100 basic solution can be used when stabilised and unstabilised. Technovit 9100 PMMA Powder - Component 2 SDS The PMMA powder is used to guarantee a clear decrease in polymerisation shrinkage, a reduction in the polymerisation heat released and a better polymerisation process. Technovit 9100 Hardener 1 - Component 3 SDS Hardening powder 1 is a peroxide compound that starts polymerisation with hardener 2. Technovit 9100 Hardener 2 - Component 4 SDS Hardening liquid 2 acts as a catalyst for hardener 1 to facilitate targeted polymerisation even at very low temperatures [< 0°C]. Technovit 9100 Regulator - Component 5 SDS This is comprised of a reactive organic compound that facilitates a regulated polymerisation with controlled low temperature spikes even for large quantities of polymerisation. PMMA-Granulate, EXART SDS This granulate acts as an additional internal filler when larger amounts (500-1000 ml) of polymer are to be used, for example, in the case of femur shaft with noncemented endoprosthses. The amount of monomer (basic solution) is thereby reduced, at the same time making the polymerization easier to control. Designation Quantity Component Number Technovit 9100 Basic Solution Stabilised 1 x 1000ml 1 Technovit 9100 PMMA Powder 120g 2 Technovit 9100 Hardener 1 8 bags, each 1g 3 Technovit 9100 Hardener 2 10ml 4 Technovit 9100 Regulator 5ml 5 Application: Fixation - tissue pre-treatment Fixation is done for 12 to 24 hours in various fixation solutions depending on the size of the tissue and the antigen/enzyme to be detected. Over fixation must always be avoided. The following fixation methods are possible for detecting antigens/enzymes: 4% neutral buffered formalin solution (0.1M phosphate or 0.02M phosphate buffer for iliac crest biopsies) 10% buffered formalin solution (0.1 M phosphate buffer) Fixation solution in accordance with Schaffer(formol/alcohol) 1.4% paraformaldehyde solution, cold (+4 to +8°C) for 24 - 28 hours (sensitive enzyme detection such as alkaline phosphatase, fixation-sensitive antigens) Dehydration, intermedium and immersion (pre-infiltration 1-3, infiltration)
Technovit H7100/H8100
Staining Protocol for Lymphoid Tissue Normal immunostaining procedures for routine markers on sections obtained from lymphoid tissue. Procedure Dry the sections for two hours at 37°C on a slide warmer. This must be done whether the slides have been stored at 4°C for some time or just collected. Pretreat the sections with trypsin at 37°C. Trypsin concentrations must be determined separately for each antibody. Use trypsin solution that has been preheated at 37°C for 30 minutes. Cover section with at least 100 micro litres of solution. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times. Preincubate in normal serum from the animal species in which the second antibody is raised for 30 minutes at 37°C. Only perform this step if aspecific background is present using a particular antibody. Drip off excess serum and apply the first antibody in an appropriate concentration and incubate for two hours at 37°C. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times. Block endogenous preoxidase in a solution of 0.05% hydrogen peroxide in phosphate buffered saline, pH 7.4, for 30 minutes at room temperature. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times. Incubate in appropriate dilutions of the secondary antibody, containing 5% normal serum for 60 minutes at room temperature. Wash in PBS for 10 minutes at room temperature. Refresh the buffer four to five times. Develop the peroxidase activity in daminobenzidine (DAB). Counterstain the sections in Haematoxylin or if a more advanced morphological detail is necessary, in periodic‐acid‐Schiff reagent. Cover with Glycerin‐gelatin and cover glass. Sectioning and Mounting Protocol Sectioning Sectioning is best done with a rotary or sledge microtome such as the JB‐4 Microtome. The features to look for are retraction of the specimen on the return stroke and motorised motion of the sample relative to the knife edge. Retraction is needed to keep the hard sample from brushing against the knife edge on the return stroke, damaging the edge and block face. Motorised sample movement is desirable to get more reproducible cutting speed and force. The best sections are obtained on these microtome with a glass knife. Glass knife strips are available from Electron Microscopy Sciences. Dry block faces are used; in fact, moisture and humidity will soften GMA blocks sufficiently to make sectioning impossible. If the block is too soft to section dry the block in a warm oven or move to a less humid work environment. Sections are collected from the knife edge using tweezers and stretched by floating on a water bath. Two factors are important when preparing the water bath: cleanliness and temperature. Any residue of soap or oil will decrease the surface tension of the water dramatically reducing its stretching ability and/or causing the section to sink. Increasing temperature also reduces the stretching ability of the bath so that at 20°C stretching of 10‐13% is possible while at 60°C stretching of 7‐9% is possible. Section stretching allows recovery of almost all the compression caused during sectioning. Properly stretched, much less than 1% of the compression in vertical dimension remains in the sections. Most procedures call for mounting your sections on slides before staining. The exception may be when staining sawn sections since these retain enough strength to be held in tweezers and hand‐dipped in the various staining solution (followed directly by coverslipping to slides). Mounting thinner sections to slides before staining also helps to prevent folds from developing in the sections. If you are using slides precoated with silane, skip the protocol for coating your own. Preparing Silanised Slides Procedure Dip slides in 2% silane/acetone solution for one minute. Dip slides in 100% acetone for one minute. Dip slides in double distilled water for one minute. Repeat step 3 with agitation. Air dry. Apply sections on a drop of water on the slide. Dry on 60°C hot plate for two to five minutes. Air dry at room temperature overnight. Dip your coated slide into the waterbath under the section and lift the section off the water surface. Dry the slides for about 16 minutes on a 60°C slide warmer. Prussian Blue Stain for Glycol Methacrylate Sections Procedure Stain in Potassium Ferricyanide solution at 60°C for 15 minutes. Filter solution after heating and before use. Wash in distilled water. Stain in Safranin O for two to five minutes. Wash in 1% acetic acid. Dehydrate in 96% alcohol two times then 100% alcohol. Clear in xylene and mount. Results Nuclei red Hemosiderin blue / green Solutions Potassium Ferricyanide Solution: Potassium Ferricyanide ‐ 1g Distilled water ‐ 50ml 2% Hydrochloric acid ‐ 50ml Safranin O: Safranin O (C.I. #50240) ‐ 0.2g 1% acetic Acid ‐ 100ml Citation Gerrits, P.O. and Smid, L., "Staining Procedures for Tissues Embedded in 2‐Hydroxyethyl Methacrylate", Heraeus Kulzer Periodic Acid Schiff Stain for Glycol Methacrylate SectionsProcedure Oxidise sections in 0.4% periodic acid for 30 minutes at 57°C. Wash in running tap water or sodium sulphite solution (to avoid a general pink background). Rinse in three times in distilled water. Schiff's Reagent for 15 minutes. Wash well in running tap water. Rinse in distilled water. Counterstain with Gill's Haematoxylin for 10 minutes. Wash well in running tap water. Dehydrate using 95% and 100% ethanol. Clear in xylene and cover slip. Results Nuclei blue Glycogen violet / red Basement membranes violet / red Mucine violet / red Solutions Gill's Haematoxylin: Haematoxylin (C.I. 75290) ‐ 6g Sodium Iodate ‐ 0.6gm Aluminium Sulphate ‐ 52.8g Distilled water ‐ 690ml Ethylene Glycol ‐ 250ml Glacial acetic acid ‐ 60ml Schiff's Reagent: Mix 0.5g of Pararosaniline (C.I.#42500) into 15mL of 1N Hydrochloric Acid. Mix 0.5g of Potassium Metabisulphite into 85ml of distilled water. Add the second solution to the first and store for 24 hours in a dark place. Bleach the resulting solution with 200mg of bone charcoal and filter. Store the colourless solution at 4°C. Sodium Sulphite Solution: 0.5g of sodium sulphite 100ml of distilled water Haematoxylin and Eosin Stain for Glycol Methacrylate Sections Haematoxylin and Eosin stain is a good general stain for many types of tissue. Solutions Gills Haematoxylin 6g Haematoxylin (C.I. 75290) 0.6g Sodium Iodate 52.8g Aluminium Sulphate 690ml Distilled Water 250ml Ethylene glycol 60ml Glacial acetic Acid Eosin 0.5g Eosin Y (Alcoholic C.I. 45380) 100ml Ethanol 96% 2 drops Glacial Acetic Acid Acid Ethanol 4ml 0f 25% HCl 100ml of 70% Ethanol Procedure Stain in Gill's Haematoxylin for 15 minutes. Wash in tap water for 10 minutes. Decolorize plastic, if necessary, with acid alcohol followed by tap water rinse. Rinse in distilled water. Scott's Tap Water for one to two minutes. Two changes of distilled water. Counterstain with Eosin for two to five minutes. Dehydrate using 96% and 100% ethanol. Clear in xylene and cover slip. Results Nuclei blue Basophilic cytoplasm blue Acidophilic cytoplasm pink Muscle tissue pink Connective tissue pink Glycol Methacrylate Embedding for Materials Samples Embedding in plastic often supplies the support necessary to successfully section various materials samples, especially prousand inhomogeous samples. GMA has been used to embed and section polymer resin, multilayer foils and films, paper, textiles and coatings in addition to a wide variety of biological material. The ability of this low viscosity resin to infiltrate effectively can make sectioning non‐biological samples fast and economical. Preparation of this type of sample is slightly different than preparing biological samples. Procedure A fixation step is, of course, not necessary for materials samples. Dehydration for GMA embedments does not have to be complete because of GMA's miscibility in water. Only if the sample is saturated with water should a schedule of increasing alcohol concentration be used. This will displace most of the water with a solvent that can be replaced with plastic. Use schedule as follows: 70% ethanol for 10minutes 96% ethanol for 10 minutes 96% ethanol for 10 minutes Absolute ethanol for 10 minutes Infiltration displaces the dehydration solvent with monomer prior to beginning the polymerisation reaction. If you have a porous sample, infiltration may be needed. Nonporous samples can skip this step. Make up your infiltrating solution according to the directions from your kit. While complete elimination of air is not needed, sealing the moulds with paraffin or plastic wrap is often done. Leave the samples at room temperature for one hour then at 37°C for one hour to complete polymerisation. If your embedments cannot be mounted directly to the microtome, an adapter must be attached. Place the suitable Technovit 3040 is an inexpensive media prepared by mixing three parts of the dry component to one part of the liquid component. Pour it into the adapter and polymerization will be complete with 10 minutes. The adapters will be firmly attached through a copolymerization of the Technovit 3040 with the GMA. You are now ready to begin sectioning with glass. Giemsa Stain For Glycol Methacrylate Sections This Giemsa stain works well for a variety of tissues. Procedure Stain in Giemsa working solution at room temperature for 1½ hours. Rinse in dilute acetic acid (four drops in 100mL DI water) for two seconds. Dip in 96% alcohol. Dip in 96% alcohol. Rinse in isopropyl alcohol three times for two minutes each. Clear in xylene and mount. Results Nuclei violet Cytoplasm blue Erythrocyten pink Solutions Giemsa Solution: 0.15g Azure A 0.30g Methylene Blue 0.36g Eosin Y 0.04g Phloxine B 50ml Glycerine 40ml Methanol Add dyes to Glycerine and methanol, mix well. Incubate overnight in 55‐58°C oven stirring occasionally. Stable for year. Giemsa Working Solution: 4mL Giemsa stock solution 40mL phosphate buffer solution Phosphate Buffer Solution: 7.25g Sodium phosphate, monobasic 2.75g Sodium phosphate, dibasic 1000ml Distilled water 10ml 10% Triton X‐100 Store at 4°C. Glycol Methacrylate Embedding for Soft Tissues Embedding in plastic provides many advantages to the histotechnologist. Thinner sections can be made providing improved detail. Better support is given to cellular components, offering improved morphology. Short, straightforward protocols are available giving minimum processing times. Improved chemistry provided by kits gives uniform results and a wider range of application. While this protocol will help get you started with embedding your soft biological samples in the Technovit GMA kits sold by Electron Microscopy Sciences, it follows the general guidelines for embedding in any source of GMA. Procedure Fixation of the biological samples can be done in any way appropriate for the work you are doing. Immersion or perfusion with 4% neutralised formaldehyde, prepared from paraformaldehyde according to Karnovsky, is usually preferred. Try to keep the sample size small using 10mm x10mm 2xmm as a maximum. Dehydration for GMA embedments does not have to be complete because of GMA's miscibility in water. Use a schedule of increasing alcohol concentration at room temperature as follows: 70% ethanol for two hours 96% ethanol for two hours 96% ethanol for two hours Absolute ethanol for one hour A short defatting step can help with infiltration. If desired, submerge the sample in acetone for 10 minutes. Make your infiltrating solution from: 100ml of Technovit 7100 resin, i.e. 2‐hydroxyethyl methacrylate (GMA) 1g of Hardener I (benzoyl peroxide) Mix using a magnetic stirrer until the benzoyl peroxide is complete dissolved. Store at 4°C in a dark bottle for up to two months. Infiltrate in a 50/50 mixture of 100% ethanol and infiltration solution for two hours. Leave the sample in 100% infiltration solution overnight. Infiltration with mild agitation and/or vacuum will be more complete and larger samples should have more infiltration steps over a longer period of time. Make up your embedding solution from: 15 parts of infiltrating solution 1 part of Hardener II Mix for one minute using a magnetic stirrer. Use the solution within 10 minutes, before polymerisation occurs. Glycol Methacrylate Embedding For Immunohistochemistry Embedding in plastic provides many advantages to the histotechnologist. Thinner sections can be made providing improved detail. Better support is given to cellular components, offering improved morphology. Short, straight forward protocols are available giving minimum processing times. In addition, improved chemistry and protocols are now available that are more antigen‐friendly. This protocol will help get you started with embedding your soft biological samples in Technovit 8100, a GMA kit sold by Electron Microscopy Sciences. Note the areas in which it deviates from the guidelines for embedding in GMA for morphology only. Procedure Use gentle agitation during fixation, washing, dehydration and infiltration for best results. Fixation of the biological samples can be done in any way appropriate for the antigen you are seeking. Cold acetone, periodate/lysine/paraformaldehyde, and buffered paraformaldehyde solutions have all been used successfully. Try to keep the sample size small using 10mmx10mm2mm as a maximum. Fixation should be done for several hours at 4°C. Wash out the fixative with 6% sucrose solution at 4°C overnight. Dehydration for GMA embedments does not have to be complete because of GMA's miscibility in water. Use acetone for one hour at 4°C changing the acetone at the beginning until it remains clear. Make up your infiltration solution from: 100mL of Technovit 8100 resin, i.e. 2‐hydroxyethyl Methacrylate (GMA) 0.6gm of Hardener I (benzoyl peroxide) Mix using a magnetic stirrer until the benzoyl peroxide is complete dissolved. Store at 4°C in a dark bottle for up to one month. Infiltrate for six to ten hours at 4°C using mild agitation. Vacuum will make infiltration more complete and larger samples should have more infiltration steps over a longer period of time. Make up your embedding solution from: 30mL of infiltrating solution 1mL of Hardener II Mix for one minute using a magnetic stirrer. Use the solution within 10 minutes, before polymerisation occurs. Cover your samples in the embedding moulds with this solution. Moulds of polyethylene such as the JB‐4 mould or Peel‐away moulds are most often used. Block holder/adapter in the recess on the mould and add mounting plastic. Alkaline Phosphatase for Glycol Methacrylate Sections Procedure Incubate sections in the incubating medium at room temperature for one to three hours. Two hours is sufficient in most cases. Wash in distilled water for two minutes. Counterstain with Nuclear Fast Red for five to ten minutes. Wash in distilled water for two minutes. Air dry and coverslip. Results Nuclei red Sites of enzyme activity blue To preserve the reaction product, selection of the right mounting (coverslip) medium is important. Solutions Incubating Medium: 5mg Naphtol As‐MX phosphate, di‐sodium salt (sigma) 0.25ml N,N‐dimethylformamide 30mg Fast blue BB (sigma) 25ml Distilled water 25ml Buffer Solution 2 drops 10% Magnesium Sulphate Solution Prepare fresh, shake well and filter before use. Buffer Solution: 2.4g 0.2M Tris (Hydroxyethyl)‐aminomethane 100ml Distilled water Adjust the pH of the buffer to 8.9 with dilute HCl and store at 4°C. Nuclear Fast Red: To 0.2g of Nuclear Fast Red add 200ml of boiling 0.5% aluminium sulphate solution Keep boiling for five to ten minutes. Allow to cool and filter before use. Acid Phosphatase for Glycol Methacrylate Section Procedure Incubate sections in the incubating medium at 37°C for five to 12 hours. Long incubation periods are needed to get significantly visible reaction product. Wash in distilled water for two minutes. Counterstain with Methyl Green for five minutes. Wash in distilled water for two minutes. Air dry and cover slip. Results Nuclei dark green Cytoplasm light green Sites of enzyme activity red Solutions Incubating Medium: Combine 20ml of buffer solution, 48ml of distilled water and 4ml of substrate solution. Combine 3.2ml of Pararosaniline solution with 3.2ml of sodium nitrite solution. Mix for one minute. Add the second solution to the first. Adjust pH to 5. Buffer Solution: 5.9 g Anhydrous sodium acetate 14.7g Sodium barbiturate 500ml Distilled water (boiled) Do not adjust the pH of the buffer and store at 4°C. Substrate solution: 40mg Naphtol As‐BI phosphatease, sodium salt 4ml N.N‐dimethylformamide Pararosaniline Solution: 2g Pararosaniline (C.I.#42500) 50ml 2N HCl Use heat to dissolve, filter when cool and store at 4°C Sodium Nitrite Solution: Sodium Nitrite ‐ 1g Distilled Water ‐ 25ml Prepare fresh and store at 4°C. Methyl Green Methyl Green (C.I.# 42585) ‐ 1g Phosphate/citrate buffer 0.1M pH 4.0 ‐ 100ml Citation: Gerrits, P. O. and Smid, L., "Staining Procedures for Tissues Embedded in 2‐Hydroxyethyl Methacrylate", Heraeus Kulzer.
Technovit® Glycol Methacrylate Embedding Kit
Technovit® 8100 Glycol Methacrylate is a HEMA-based plastic-embedding system for studies with light microscopy. It is suitable for embedding all tissues in medicine, zoology and botany. Sections of decalcified or briefly decalcified iliac crest biopsies and implanted biomaterials can be used for more than just histological staining; they can also be used for enzyme chemistry and immunohistochemistry. Material properties Technovit® 8100 is a combination of a practically odourless plasticiser and a hydrophillic plastic. Technovit® 8100 was specifically developed for cold polymerization (+4°C). Note: While hardening the embedding form must be hermetically sealed because the polymerisation system is oxygen-sensitive. Overview of the benefits Reproducibility and reliability of the embedding due to the constant, documented quality controls of the individual components Low polymerisation temperature of 10°C to 0°C due to the special catalyst system and the PTFE forms Uniform hardening of the block, thus uniform and thinnest possible sections Low shrinkage artefacts, thus excellent tissue morphology Routine staining, enzyme detection and immunohistochemistry possible Haemotological illiac crest biopsies do not need to be decalcified Low toxicity due to special combination of plasticiser and catalyst system Application Prepare Technovit® 8100 in accordance with the step-by-step instructions. Place the fixated and dehydrated specimens in the infiltration solution. A low temperature and agitation of the specimens is beneficial during the entire embedding process. Polymerisation Prepare the polymerisation mixture according to the instructions and then fill the embedding cavities. Position the infiltrated specimens therein. Hermetically cover the cavities with films. Place on a pre-cooled gel plate or thin layer of ice at 4°C to harden. The films are removed after polymerisation is complete and blocked with Histobloc® and Technovit® 3040. It is not possible to elute the plastic before staining or reaction. Product data Designation Quantity Technovit® 8100 1 x 500ml Basic Solution 5 x 0.6g hardener 1 1 x 30g hardener 2 1 x 500pcs PE films Technical data Colour Transparent Density = spec. weight g/cm3 (DIN 53479) 1.08 Refractive index Monomer Polymer 1,4485 1,4990 Storage temperature max. 25°C Shelf life 2 years The following instructions for fixation and dehydration are not necessarily required. Technovit® can also be infiltrated and polymerised after other pre-treatment. Airtight glass or PE disposable containers (approx. 20ml) must be used for the entire process! Tip: The specimens must be constantly agitated during fixation, dehydration and infiltration! Fixation In order to achieve optimal immunohistochemical results, it is recommended to work at 4°C throughout the entire embedding process and to aim for short fixation times. Fix the smallest possible pieces of tissue (1mm thickness) in 2% paraformaldehyde in phosphate buffer pH 7.4 at 4°C for 3-4 hours. Subsequently, retreat for 12 hours (overnight) in phosphate buffer pH7.4 with an additional 6.8% sucrose at 4°C. Dehydration Dehydrate the tissue in cold acetone 100% for at least one hour at 4°C. Change as often as possible in the first minutes until the acetone remains clear. Infiltration Making the infiltration solution: Technovit® 8100 Basic Solution 100ml + Technovit® 8100 hardener 1 1 bag, 0.6g Dissolve in a clean, detergent-free PE or glass container and then place at 4°C. When sealed, the infiltration solution is stable for a maximum of four weeks at 4°C. Transfer the specimen directly from the acetone to the pre-cooled infiltration solution. The specimens remain therein for 6-10 hours at 4°C. Polymerisation Making the Polymerisation solution: Infiltration solution, 4°C 15ml + Technovit® 8100 hardener 2 cooled 0.5ml Measure with standard pipetting aids and mix well in a PE or glass container. Then, carefully mix the infiltrated specimen in a sealed container for approx. five minutes. The colour of the polymerisation solution changes first to yellow-green, but after hardening it becomes colourless. Completely fill the Histoform cavities with a disposable pipette, position the tissue therein and immediately cover with transparent PE film. Multiple films can be used for a cavity in order to hermetically seal the cavity. Do not press out bubbles; rather, apply more polymerisation solution and add new film. During polymerisation (at least 3 hours) the embedding form must be placed on a cooling plate or thin layer of ice at 4°C. Do not let the form or specimens come into contact with moisture. Histoform Q Material Room temp. approx. +20°C Refrigerator +4°C Refrigerator on ice 0°C Technovit® 8100 30:1 -- 69 48 Technovit® 8100 35:1 -- 52 42 Technovit® 8100 40:1 -- 50 41 Histoform S Material Room temp. approx. +20°C Refrigerator +4°C Refrigerator on ice 0°C Technovit® 8100 30:1 69 21 12 Technovit® 8100 35:1 67 19 11 Technovit® 8100 40:1 65 -- -- Blocking and archiving Remove the film at room temperature with tweezers once hardening is complete. The specimens are blocked with Histobloc® and Technovit® 3040 so that they can be removed from the PTFE mould. Store blocks that are not needed immediately (for immunohistochemistry) at a cool temperature in plastic bags or similar. Processing One obtains the best cutting results with a rotation microtome, with the Technovit® Histoblade in combination with the Heraeus knife holder or a hard metal knife (glass diamond knife). Tightly clamp the blocks in the totem cam system on the microtome. Dryly remove the sections with forceps and place in a bath (Aqua dest.). Place directly on a coated object holder and let dry for 2 hours or more at 37°C. Dry sections that are not needed immediately (for immunohistochemistry) at room temperature and store for a maximum of five days at 4°C. Polymerisation 15ml, 5ml at 4°C Overview of how to make the solution Solution Ethanol Basic solution Technovit® 8100 Hardener 1 Technovit® 8100 Infiltration Solution Hardener 2 Technovit® 8100 Application-temp Infiltration 100ml 0.6 (1 bag) 4°C Object holder coating For example, submerse the object holder in a solution of 0.5% Alcian blue (8GXL Sigma) at 65°C for 15 minutes or liquidly coat the object holder with 0.1% poly-L-lysine (Sigma). All standard coated object holders may be used. The sections must dry for at least two hours at 37°C. Place the non-deplasticied sections directly in the stain solution or start with enzymatic pre-treatment. Example Enzymatic pre-treatment: Incubate sections for 5-10 minutes in 0.01% trypsin with 0.1% CaCl (calcium chloride) pH 7.8 Wash multiple times in phosphate buffer (PBS) for five minutes Incubate for two hours at 37°C with primary antibody, change multiple times Block the endogenous peroxidase with 0.06% hydrogen peroxyde in phosphate buffer (PBS) (30 minutes at room temperature) Wash multiple times in phosphate buffer (PBS) for five minutes Incubate with the second antibody for 30 minutes at room temperature Wash multiple times in phosphate buffer (PBS) for five minutes Diaminobenzidine (DAB) as for cyrostat sections 10-15 seconds of counterstaining with hematoxylin Blue for three minutes under flowing water Cover with glycerine gelatine Determination of immuno factors is possible with AP, PAAP, APAAP, ABC, avidin-biotin, streptavidin and immunofluorescence methods. The use of wetting agent, e.g. Tween, in the rinsing buffer is discouraged. The peroxidase should be dissolved in buffer. With the constantly changing range of new products for histochemistry and immunohistochemistry, it is always advisable to follow the respective manufacturer instructions.
Terg-A-Zyme®
Terg-a-Zyme consists primarily of a homogeneous blend of sodium linear alkylaryl sulfonate, phosphates, carbonates, and protease enzyme. Terg-a-Zyme is anionic in nature. The protease enzyme in Terg-a-Zyme is bacillus licheniformis subtilisin carlsberg, which may be deactivated by 300ppm hypochlorite at 30°C in seconds; 3.5ppm hypochlorite at 38°C for 2min; exposure to pH below 4 for 30min at 60°C; or by heating to 80°C for 10min. Why Terg-A-Zyme® Terg-a-Zyme is an enzyme-active powdered detergent that is versatile in that it Is concentrated to save you money Is biodegradable Replaces corrosive acids and hazardous solvents Has protease enzyme that remove proteinaceous soils, tissue, blood, and body fluids Is of free-rinsing to give you quality results Is used to pass your cleaning and inspection approvals Terg-a-Zyme is used to clean Hospital instruments Dairy equipment Laboratory ware Reverse osmosis Ultrafiltration membranes and units Sampling apparatus Pharmaceutical apparatus Cosmetics manufacturing equipment Tubing Pipes Optical parts Process equipment Industrial parts Desalination plants Terg-a-Zyme is used to remove Soil Grit Grime Blood Tissue Grease Fat and oils Proteinaceous soils Dairy proteins Particulates Solvents and Bioreactor residue Terg-a-Zyme cleans a number of surfaces, such as Glass Metal Stainless steel Porcelain Ceramic Plastic Rubber Fiberglass Soft metals Copper Aluminum Zinc Magnesium Terg-a-Zyme should follow cleaning methods Soak Brush Sponge Cloth Ultrasonic Flow through clean-in-place Will foam – not for spray or machine use Directions Make a fresh 1% solution (10g/L) in cold or warm water. If available, use warm water below 55°C. Clean by soak, circulate, wipe, or ultrasonic methods. Follow manufacturer's directions for filter membrane cleaning. Rinse thoroughly, preferably with running water. For critical cleaning, do final or all rinsing in distilled, deionised, or purified water. For food contact surfaces, rinse with potable water. Cleaning Validation Methods Test a parameter of rinse water before and after rinsing the cleaned surface, or test the clean surface. No significant change in the parameter indicates no detectable detergent residue. Parameters measured include: pH, conductivity, UV, TOC, HPLC, sodium concentration, phosphorus concentration, anionic detergent concentration using inexpensive detergent water testing kits, surface tension, and surface analysis. Technical Data Physical Data Typical Value pH of 1% solution 9.5 Flash point (°C) None Phosphate content (as phosphorous) 7.5% Organic carbon (1% calculated w/w) 11% Fragrance content 0% Surface tension 1% solution (Dyne/cm) 32 Percent active ingredients 100% Color White and cream-coloured flakes and brown specks Form Powder Solubility in water To 10% (w/w) at ambient temperature Hard water effectiveness Highly effective Biodegradability Biodegradable Foam tendency High-foaming Shelf life Two (2) years from the date of manufacture
Tissue Grip Water Bath Adhesive
Introduction Liquid water bath adhesive is used in Histology & Embedding staining, special stains, and immunoperoxidase procedures. Uses/Limitations For in-vitro diagnostic use only Do not use if reagent is cloudy Do not use past expiration date Procedures Fill the water bath with water (DI or tap) and add 5-10ml per liter of warm water (40-46°C) to the bath. Stir the water gently to ensure Tissue Grip dissolves completely and is mixed well. Follow Lab SOP for section recovery, drying, and staining. Special stains and immunohistochemistry may be performed on sections, recovered with Tissue Grip™. Avoid the use of adhesive, coated or charged slides. For frozen sections, acetone cleaned slides may be dipped into a 1:10 solution with water, allowed to dry, and stored in a slide box for later use. Clean flotation bath after use. Storage Storage: room temperature. The product may become semi-solid when exposed to colder temperatures. If the solution has become solidified, move to a warmer area and allow it to "thaw" for 24-48 hours.
Tissue-Tack Adhesive
PRODUCT DISCONTINUED BY SUPPLIER BACKGROUND: Loss of sections during lengthy histochemical procedures causes frustration and diagnostic delays. Tissue Tack is an acrylic-based adhesive which has been used successfully to attach JB-4 sections to slides when conventional section adhesives have proved to be inefficient. Application of Tissue Tack in the following method allows for the effective demonstration of immunofluorescent and immunoperoxidase staining. This adhesive is also applicable to the Esterbauer and Nohammer modification of the DDD test. Dry microtomed JB-4 sections onto a clean glass slide. A small perforation should be made in the JB-4 plastic around the perimeter of the tissue section, to expose the glass. Searing the edges of the plastic with a hot instrument provides additional mechanical adhesion. Using an applicator stick, apply a small amount of Tissue Tack, covering the perforated area of the JB-4 section. Quickly smear the adhesive in the direction away from the section to form a thin film over the slide. (This procedure should be performed under a fume hood.) Air dry for a few minutes in the fume hood, until the adhesive is clear and not sticky. Proceed with desired procedures. Tissue Tack becomes soft and tacky when it is subjected to strong alkalis and solvents, and adheres to the glass slide, but not to the JB-4. It can be removed after the staining techniques. CAUTION: Prolonged exposure can cause irritation to eyes, skin and respiratory tract. Wear protective glasses and gloves. Use only with adequate ventilation. Keep the container tightly closed.
Triphan Replicating Sheets
PRODUCT DISCONTINUED BY SUPPLIER (This is a direct replacement for Triafol) Electron Microscopic Specimen Preparation Triafol Replicas: Application: The Triafol technique is a method for making electron microscopic specimens which corresponds to a great extent to the indirect replica method described by Mahl and Konig. Triafol foil (trade name for acetobutyrate foil) Instead of varnish composed of pyroxylin and amyl acetate is used for taking replicas. The preparation time is thereby greatly reduced while the quality of the substances remains the same. Whereas pyroxylin-amylacetate (zaponlac) replicas may not be removed from the base until they have dried for 10 hours, the fast drying Triafol foils may be removed after only 3 to 5 minutes, without having to expect deformations. The entire preparation process from taking the replica to finished specimen requires approximately two hours. Mode of Operation: For reproducing smooth and moderately rough surfaces, foil of 0.04mm thickness is suitable while 0.1mm Triafol foil should be used for rougher surfaces. Depending on the size of the surface to be replicated, cut small foil pieces and clean carefully with a soft brush removing any adhering particles. Wet the preparation spot, which is to be replicated, with a few drops of acetone (spray from a fine glass capillary tube). If too much solvent is used, there is the danger that the thin foil expands too much and then tears when it is removed. Before the solvent can evaporate apply the Triafol foil to the specimen surface with as little pressure as possible and wet the foil with acetone. Always use the dull side of the foil for taking replicas. The dried Triafol replica can be removed from the base after 3 to 5 minutes. Coat the foil with a thin metal or metal oxide layer in an evaporation unit. After oblique shadowing (angle 15 to 25 degrees for smooth objects, 30 to 45 degrees for rough objects) and application of a very thin vertical vapourization layer, coat the foil on the vaporised side with a paraffin film of approximately 0.3 mm thickness in order to render the thin vaporisation layer more solid. This layer can easily tear during the subsequent solution process because the lacquer expands greatly when it enters into contact with a solvent. When the molten paraffin (melting point approximately 52 to 53°C) is applied from an open glass tube, make sure that no paraffin is applied to the back of the foil. Attention: For the dissolving processes described below, make sure that there is absolute freedom from vibrations because the slightest oscillation causes tearing of the thin replica foils and renders the replica useless due to the formation of folds. Place the matrix cautiously into a methyl acetate bath saturated with paraffin (paraffin side upwards) until the violet colour of the Triafol foil has disappeared. Remove the matrices after 20 minutes with a glass siphon and rinse again for 5 to 10 minutes in a second fresh methyl acetate bath. Then, dry matrices In a dust-free place on filter paper, with the shadowing layer upwards and the paraffin layer downwards. Then, cut specimens with a razor blade into small squares having an edge length of approximately 2mm; place these (vaporisation layer downwards) on the surface of clean electron microscopic specimen carriers (specimen diaphragms or grids), and weigh them by placing a small piece of brass wire mesh (0.3mm wire, 0.6mm mesh width, size 2mm x 2mm) on them. Place the specimen diaphragms, which have been weighed in this manner, in a bath with heated toluene p. A. with forceps for removing the paraffin. Leave them in the toluene bath until the paraffin has been removed completely (approximately 10 minutes). Remove all paraffin traces in a second toluene bath. Take out specimen carrier cautiously and dry carefully. The replica foil may easily tear if the liquid which is contained in that diaphragm cone is sucked out suddenly. Therefore, place the diaphragms on the filter paper first with one edge only. When the specimens are dry, the wire meshes can be thrown off easily. Contaminated or damaged specimens can usually be eliminated by light microscopic control.
Uranyl Acetate
Ordering procedures required to satisfy Federal and Queensland policies/ legislation Pricing and shipping of uranyl acetate: The pricing indicated in our online catalogue lists 2 separate items: the cost of uranyl acetate and the cost for shipping. The shipping cost applies regardless of how many units are ordered and is valid to anywhere in the world. If you fax us a copy of the permit from the radiation control authority, we will ensure that this is affixed to the order, which will be shipped direct to you. The Australian Authority may insist on sighting the original permit, but it can pass through Customs on the copy. The permit for Uranyl Acetate can be obtained from the Australian Radiation Protection and Nuclear Safety Agency (ARPANSA). Their details are: Phone: (03) 9433 2211 Fax: (03) 9432 1835 Email: regulatory.unit@health.gov.au Measurements on 100g sample of Uranyl Acetate. All measurements have been done above an open bottle. Alpha (a) using a 540 scintillation meter with AP-2 Probe Beta (b) >500 Counts/sec using a 540El probe coupled to a GM Meter (this determines beta events and some low energy gamma events. Gamma dose Rate (energy field) (g) Two measurements done: Please note that a high efficiency scintillation counter will measure many of these events, but it will say nothing about the energy field. The customer must use a dose rate meter. (You can see the difference between the energy events and gamma dose rate) A Geiger counter just will not do the job. Mini monitor type R with GM Probe - This is more specific to Gamma due in fact to being constructed to accept a narrow energy range (discriminates against beta) field: 0.6mR/hr (Mainly gamma)? Ionization chamber DMM 95/0500 - 1/6 for total Beta and Gamma energy field 5mR/hr (g b) Our uranium is U235 depleted. Typical Isotopic Composition is: U238 - 99.6 - 99.7% U235 - 0.3 - 0.4% Natural Uranium has two main Isotopes: U238 & U235 Natural Isotopic composition would be: U238 - 99.3% U235 - 0.7% That is to say material supplied by us is Uranium depleted i.e., the U235 content is reduced from 0.7% to between 0.3 - 0.4%. Both natural and depleted Uranium, being a mixture of isotopes and daughters, will be expected to demonstrate alpha, beta and gamma activity. RADIOACTIVE MATERIAL The average number of atomic transformations occurring per second is termed the activity of the radioactive material. The old, traditional, unit of activity is Curie(Ci), which is defined as 3.7 x 1010 atomic transformations per seconds. For smaller quantities the one-thousandth and one-millionth unit are used, namely the millicurie (mCi) and the microcurie (µCi). However, by International agreement, the unit of activity is currently being changed. The new unit of activity is the becquerel (Bq) which is defined as one atomic transformation per second. Becquerel is an extremely small unit. Larger units are used, namely the kilo-becquerel (1kBq = 103Bq), the mega-becquerel (1MBq = 106Bq), the giga-becquerel (1GBq = 109Bq) and the tera-becquerel (1 TBq = 1012Bq). It may be noted that: 1 Ci = 3.7 x 1010Bq = 37GBq = 0.037TBq. A material must have a specific activity greater than 74 becquerel per gram (Bq/g) (0.002µCi/g) or 74 kBq/kg in order to be regarded as a radioactive material. Uranyl Acetate sold by ProSciTech Pty. Ltd. has an activity of: U (depleted) .51µCi·gm-1 (µCi = microcuries) Formula for conversion to Bq is: 1 Ci = 3.7 x 1010Bq ?\ for U (depleted) Specific Activity = 0.51 x 10-6 x 3.7 x 1010Bq·gm-1 = 1.887 x 104Bq·gm-1 Uranyl Acetate Specific Activity (U approx. 55%) = 1.04 x104Bq·gm-1 \ Uranyl Acetate will exhibit 10, 400 disintegrations/sec/gm.
Uranyl Acetate Replacement Stain (UAR-EMS)
EMS22405 Description UAR-EMS Stain is a new negative and positive stain which may be used as an alternative to uranyl acetate. Procedure The UAR Stain is supplied as a concentrate. Please use the following dilutions: For most very small particulate specimens like most viruses, dilute by 4x with distilled water. If you are working with bacteria, then a dilution of 15-40x is needed. Use similar dilutions for positive and negative staining. Positive staining will take place in 30 minutes at room temperature. For increased contrast of the stain, simply adjust the staining time. Reference Article Nakakoshi, Masamichi, Hideo Nishioka and Eisaku Katayama, 2011. New versatile reagents for biological transmission electron microscopy that substitute for uranyl acetate. Journal of Electron Microscopy, 60(6): 401-407. Abstract Aqueous uranyl acetate has been extensively used as a superb staining reagent for transmission electron microscopy of biological materials. However, recent regulation of nuclear fuel material severely restricts its use even for purely scientific purposes. Since uranyl salts are hazardous due to biological toxicity and remaining radioactivity, development of safe and non-radioactive substitutes is greatly anticipated. We examined two lanthanide salts, samarium triacetate and gadolinium triacetate, and found that 1-10% solution of these reagents was safe but still possess excellent capability for staining thin sections of plastic-embedded materials of animal and plant origin. Although post-fixation with osmium tetroxide was essential for high-contrast staining, post-staining with lead citrate could be eliminated if a slow-scan CCD camera is available for observation. These lanthanide salts can also be utilised as good negative-staining reagents to study supra-molecular architecture of biological macro-molecules. They were not as effective as a fixative of protein assembly, reflecting the non-hazardous nature of the reagents.