0.  Safety precautions

  • You must complete required lab safety training before starting this procedure.
  • If this is your first time doing this procedure, ask to be trained by an experienced lab member.  If you have not done this in a while, you should ask for a refresher.
  • Before starting, even if you have done this procedure before,
    • read this protocol entirely
    • review relevant Safety Data Sheets and Harris Lab SOP (also see below)
    • ensure you have all reagents and supplies listed below
    • ensure all equipment is in good working order
    • have all waste containers ready (also see Clean-up).
    • plan your schedule well so that you wouldn’t have to rush
  • Review SDS and Harris Lab SOP for the following hazardous chemicals used in this procedure:
    • 1,2-dichloroethane: flammable, acute toxicity (oral & inhalation), irritation (skin & eye), carcinogen
    • Ethanol: flammable, irritation (eye)
    • 1,2-dichloroethane and PEI solution must be handled only under a chemical fume hood, with double-layer of nitrile gloves on.
  • The following Personal Protective Equipment is required for this procedure:
    • Lab coat
    • Nitrile gloves (double-layer required; regularly check for holes)Eye goggles
  • Place a piece of absorbent sheet on the work surface before starting the procedure.

1.  Reagents and supplies

1.1  For cleaning grids

  • Grids (SynapTek Be-Cu Notch-Dot 1 × 2 mm slot; Ted Pella 4516)
  • 20-ml borosilicate glass scintillation vial × 1
  • Sonicator (stored under a fume hood in NHB 3.360E)
  • Water (RO or deionized)
  • Kimwipe
  • disposable pipets
  • Ethanol (100%; EMS 15056; use excess from epoxy embedding)
  • 1,2-dichloroethane (Sigma-Aldrich 284505 or Fisher E175-500)
  • 90-mm glass Petri dish lined with filter paper
  • Foreceps

1.2  For PEI solution

  • Polyetherimide (PEI; Goodfellow 292-028-58)
  • 1,2-dichloroethane (Sigma-Aldrich 284505 or Fisher E175-500)
  • 250-ml Pyrex glass bottle × 1
  • 10-ml borosilicate glass pipette
  • Scale (2000 g max.; in a fume hood in NHB 3.360H)
  • Aluminum foil

1.3  For grid coating

  • 250-ml glass beaker (dedicated for use with 1,2-dichloroethane) × 1
  • Funnel support
  • Glass funnel, 125 mm top diameter (e.g., VWR 89000-472) × 1
  • Glass Coplin jars × 3; one to be used for PEI solution should have a screw-cap with foil lining
  • Millipore Swinnex 47 mm Filter Holder (EMD Millipore SX0004700 [Fisher SX0004700])
  • 50-ml glass syringe (e.g., Cadence Science 5059 [Fisher 14-825-11C])
  • Glass microscope slides (Gold Seal Microslides, Fisher 12-518-101)
  • Sonicator (stored in NHB 3.360, next to sink)
  • Lens tissue (EMS 71700)
  • Desiccator jar (filled with silica gel desiccant) with vaccum tubing attached
  • Microscope slide grip (optional)
  • Whatman Grade 1 Filter Paper, 240 mm diameter (Whatman 1001240 [Fisher 09-805J])
  • Molecular Sieve, Type 4A, 8-12 Mesh Beads (Fisher M514-500)
  • Glass jar with lid (filled with silica gel desiccant [e.g., Ted Pella 19961] and lined with several sheets of filter paper)
  • Millipore Nylon hydrophilic membrane filters, 47 mm diameter, 0.2 µm pore (EMD Millipore GNWP04700 [Fisher])
  • Support clamp
  • Plexiglas box stage
  • Lamp
  • Pyrex glass crystallizing dish, 190 mm diameter × 100 mm height (Corning 3140190 [Fisher 08-741H]), covered with matte black paper
  • Single-edge razor blades
  • Parafilm, cut into 2-inch squares
  • 150-mm plastic petri dish, lined with filter paper
  • Grid coating pen (EMS 70624)

2.  Procedures

2.1  Cleaning grids (should be done the day before use or earlier)

  • Perform this procedure in a fume hood lined with absorbent sheet.  Wear two layers of nitrile gloves, lab coat, and eye protection.
  • Follow this procedure to (1) prepare new grids, or (2) reuse grids previously coated with PEI (either failed quality checks or hole in pick-up; discard grids that are overly oxidized or bent).
  1. Place grids into a 20-ml scintillation vial.
  2. Add ~10 ml of 100% ethanol into vial.
  3. Loosely cap the vial and place it into sonicator (located in the cabinet at the lower fume hood) filled with RO water (water level should be about the same as ethanol in the vial). 
  4. Sonicate for 5-10 min. 
  5. Take the vial out of sonicator and wipe water off with Kimwipe.  Use a disposable pipet to remove ethanol. Collect the waste ethanol into "Flammable" waste bottle (located in flammable storage cabinet). 
  6. Repeat Steps 2 – 5.
  7. Add ~10 ml of 1,2-dichloroethane into vial and sonicate for 5-10 min. Follow Step 5 to collect waste 1,2-dichloroethane into “1,2-dichloroethane/PEI” waste bottle (located in flammable storage cabinet).
  8. Repeat cleaning with 1,2-dichloroethane two more times.
  9. After removing 1,2-dichloroethane, wipe the outside of vial with Kimwipe to remove water.  Tap out the grids onto a glass Petri dish lined with filter paper. As the grids dry out, spread them into a single layer using the back side (curved) of a pair of forceps.
  10. Keep lid of the dish ajar and let the grids air dry overnight in fume hood.
  11. Discard the used vial, pipets, and Kimwipes into “Solid Waste – No UA” waste bag. Empty the sonicator and store it in the cabinet at the lower fume hood. Return ethanol, 1,2-dichloroethane, and their waste bottles into the flammable storage cabinet.

2.2  PEI solution (should be made 72 hrs before use or earlier)

  • Perform this procedure in a fume hood.  Wear two layers of nitrile gloves, lab coat, and eye protection.
  1. Rinse a 250-ml Pyrex bottle with a small amount of 1,2-dichloroethane, pour into the appropriate waste container.  Let the bottle air-dry in a fume hood overnight.
  2. Bring a large scale (max. = 2000 g; usually located in a fume hood in NHB 3.360H) into a fume hood in NHB 3.360E and place a piece of aluminum foil on top.  Tare the air-dried Pyrex bottle.
  3. Measure out 0.7 g of polyetherimide (PEI) using scale by refrigerator in NHB 3.360E.  Add PEI to the Pyrex bottle. 
  4. Add 1,2-dichloroethane with a 10-ml borosilicate glass pipette to a total weight of 200 g. (Final PEI concentration = 0.35% [w/w])Note 1 
  5. Let the mixture stand in the flammables cabinet. Gently swirl the solution after 24 hrs.  It will take at least 48 hrs for PEI to dissolve.Note 2
  6. Return Ethanol, 1,2-dichloroethane into the flammable storage cabinet, and the scale to NHB 3.360H.

Note

1: It may be necessary to adjust the concentration of PEI solution to achieve optimal thickness (see “Tips and troubleshooting”).

2: Do not cast film unless PEI is completely dissolved. "Fresh" PEI solution still contains long strands of undissolved polymer and does not form consistent thin films. Occasional swirling after 24 hrs helps PEI to dissolve completely. Suggested schedule: mix PEI and 1,2-dichloroethane on Thursday, swirl the solution just before you leave on Friday, then let it stand over the weekend.

2.3  Cleaning equipment (should be done the day before use or earlier)

  • Perform this procedure in a fume hood. Wear two layers of nitrile gloves, lab coat, and eye protection.
  • Equipment to be cleaned: glass funnel, glass Coplin jar, glass syringe, Swinnex filter holder
  1. Place the funnel over a Coplin jar (with foil-lined screw-cap) using a funnel holder. Assemble the filter holder (without filter membrane) and glass syringe.  Have a piece of aluminum foil spread on work surface in a fume hood.
  2. Pour about 100 ml of 1,2-dichloroethane in a dedicated 250-ml glass beaker.
  3. Rinse the syringe by passing 1,2-dichloroethane in and out of it several times. The plunger should slide smoothly without resistance.
  4. Draw 1,2-dichloroethane into the syringe and attach filter holder. As 1,2-dichloroethane is expelled, rinse the inside of funnel. Let the rinse collect into the Coplin jar, and discard in “1,2-dichloroethane/PEI” waste bottle.
  5. Fill about half of the Coplin jar with 1,2-dichloroethane, close the cap and gently shake to rinse the jar.  Discard the rinse in the same waste bottle.
  6. Disassemble the filter holder and syringe and place on aluminum foil.
  7. Leave the rinsed equipment out to dry overnight in fume hood.
  8. Return 1,2-dichloroethane and the waste bottle into the flammable storage cabinet.

2.4  Coating

2.4.1  Preparing Glass Slides (should be done the day before use or earlier)

  • This part can be performed in the grid coating station in NHB 3.360. Wear nitrile gloves.
  1. Place glass slides into a Coplin jar (not the same one used with PEI) containing 100% ethanol and sonicate for 5-10 min.
    1. Visually inspect the glass slides for any chips and scratches. Use only the clean ones.
  2. Dry slide by absorbing as much ethanol with ~5 plies of lens tissue. Scrub the slides clean with 1 ply lens tissue at least twice. Try not to shred the tissue – it you see lint on the glass surface, use air duster to clean.
  3. Store the cleaned slides until use in another clean, dry Coplin jar, placed in a desiccator jar with vacuum on.
  4. Collect the waste ethanol into "Flammable" waste bottle. Rinse the Coplin jar with RO water and air-dry. Discard used lens tissue as general trash. Return ethanol and the waste bottle into the flammable storage cabinet.


2.4.2  Casting PEI Film

  • This part must be performed in the fume hood.  Wear two layers of nitrile gloves, lab coat, and eye protection.
  1. Assemble the filtration device:
    1. Install a piece of Millipore filter membrane into the cleaned filter holder. The holder has two gaskets – wet the inner gasket with a small amount of 1,2-dichloroethane so that it would expand enough to fit the holder. Then place a piece of filter membrane on top of the inner gasket. Screw the holder tightly enough to make a good seal, but not too tightly to break the membrane (you’d hear a crack when this happens). Attach it to glass syringe with the plunger removed.
    2. Place the syringe-filter on top of Coplin jar cleaned with 1,2-dichloroethane.  Use a clamp to support the syringe.
    3. Place glass funnel over the syringe with a funnel support.  The tip of funnel should nest inside the syringe.
    4. Place a piece of Whatman filter paper in the funnel and add some molecular sieves.
  2. Filtration of PEI solution:
    1. Add ~50 ml of the PEI solution into the funnel. The solution should collect in the syringe.
      1. If it passes rapidly into the Coplin jar, the filter membrane is cracked or not installed correctly. Place the PEI solution back to the Pyrex bottle, replace the filter, and try again.
    2. When ~35 ml (or enough to soak 2/3 of microscope slide length) of PEI solution has accumulated in the syringe, remove the funnel (place it on the PEI solution bottle), apply the plunger to filter the solution through Millipore filter into Coplin jar.
    3. Cap the Coplin jar and place in a desiccator jar filled with desiccant (but do not apply vacuum).
    4. Discard the Whatman filter paper with molecular sieves into “Solid Waste – No UA” waste bag.  Cap the PEI solution bottle.
  3. Casting PEI film:
    1. Place the Coplin jar containing filtered PEI solution in a fume hood.
    2. Dip the cleaned slide in the PEI solution, remove it quickly in a single smooth pull, and place it in a filter paper-lined glass jar filled with desiccant. This can be done one slide at a time, or as multiple slides in a slide holder all at once.
    3. Close the jar and place in a desiccator jar with vacuum on to let slides dry for at least 45 minutes.
  4. Cleaning the filtration equipment:
    1. Remove the filter membrane from the filter holder and discard into “Solid Waste – No UA” waste bag.
    2. Clean the equipment with 1,2-dichloroethane as described above.

2.4.3  Coating Grids

  1. Carry the jar/slide over to a lab bench where the Plexiglas box stage is set up.
  2. Fill a large glass container (crystallizing dish covered with matte black paper) with RO water.  Avoid touching the water with your hands, as the water must stay clean.
  3. Turn on the lamp and adjust its angle and/or location, such that the reflection on water surface can be seen easily.
  4. Remove the slide from the jar by holding the frosted end.  Brace the slide at an angle against a few layers of lens tissue on bench top.
  5. Use the corner of a new single-edge razor blade to make cuts along the edges of both sides of the glass slide. This breaks the continuous PEI film so it can be stripped off.
  6. Fog the film by breathing on it heavily. 
  7. Holding the frosted end, briefly submerge the tip of slide (about 1/8 in.) into the bowl of water (just to wet the slide).
  8. Slowly submerge the length of the slide into the water.  The film from each side of the slide should release onto the surface of the water.  If the film resists coming off the slide in any way, don't force it.  The stretching introduces artifact on the film – discard that slide and go on to the next.
  9. Adjust the lamp from above to reveal the interference color of the film – the color should be gray to silver.  Note any wrinkles and blemishes in the films in worksheet.  Look for the best areas on it to drop grids – you'll be able to see the imperfections of the film and be able to avoid the edges and end that are often distorted or thicker.  Discard any unusable films, by wiping the surface of the water with a lens tissue.
  10. With the curved forceps, place grids in a rectangular pattern on the good areas of the floating film (e.g., up to about 4 × 8 grids).  Place shiny side of the grid on the film (i.e., notch side up, if using notched Synaptek slot grids).
  11. Return to each grid and gently press one edge of the grid lightly, with a closed forceps.  This will help to attach the grid to the film.
  12. Wrap a clean dry glass microscope slide with a 2-in. square piece of Parafilm while keeping the paper backing. Press onto Parafilm or lightly scratch on the paper backing to get Parafilm stick onto the slide.  On frosted end of the slide, write down the batch information (this usually consists of today's date and which side of which slide the film was casted on; e.g., "21030126-3b", for a film casted on Jan. 26, 2013 on the backside of slide #3.).  Do not remove the paper backing until ready to pick up the Pioloform film containing grids.
  13. Have ready the plastic Petri dish sets (150-mm diameter) with filter paper lining the bottom dish.  Label the dish with today’s date and with anything else appropriate.  One dish set can accommodate up to 5 slides.
  14. Remove the paper backing of Parafilm, and draw two straight lines with a grid coating pen – one along the end of the Parafilm-covered slide, and another about 1.5 in. from it (toward the frosted side of the slide).
  15. Position the end of the Parafilm-covered slide above the “free” end of a floating film.  Then dip slide into the water to catch this free end and continue to plunge the slide deeper straight into the water in order drape the entire film of grids onto the glass slide.  Then pull the slide straight up out of the water. Invert the slide for a minute to drain any water out of the slide.  You can also flick excess water off the slide.
  16. Place the microscope slide containing the grids, grid side up, into the 150-mm plastic Petri dish for storage.  Allow the grids to air dry overnight.  For best film adhesion, put the slide with grids in the oven overnight at 60-70°C.  For best results, the grids should be used on the next day after quality check.


3.  Clean-up

3.1  Waste containers:

  • Hazardous Liquid Waste: Pour all waste into the proper waste collection bottles stored in flammable cabinet.
    • PEI-dichloroethane (1,2-dichloroethane from cleaning grids and glassware; old PEI solution)
    • Flammable Solvents (100% Ethanol used to clean grids; this ethanol should NOT contain anything else)
    • Hazardous Solid Waste: Place all contaminated solid waste (e.g., gloves, weighing dishes, filter paper/membrane, molecular sieves, aluminum foil, absorbent paperetc.) into hazardous waste bags in fume hood.
    • Sharps: Place all used razor blades and contaminated broken glassware into a sharps container.
    • Uncontaminated glass: Place all used intact glass slides into the uncontaminated glass disposal container.

3.2  Glassware and equipment:

  • Rinse glassware with 1,2-dichloroethane as described above.  Discard the rinse 1,2-dichloroethane into " PEI-dichloroethane" waste bottle.  Leave the disassembled glassware on a piece of aluminum foil in the fume hood to air dry overnight. On the next day, store the device in an appropriate area.
  • All glassware that did not come in contact with PEI solution should be rinsed with RO water and air-dried.


4.  Film quality check with tSEM

  • The main purpose of this procedure is to check the quality of newly cast PEI film at the conditions (i.e., magnification, brightness, and contrast) actually used for serial section imaging.
  1. Pick one grid per film (from near the edge of the thickest part of film) and load onto a tSEM specimen holder (i.e., “STEM 12” style).  Also load a grid containing test thin sections of good quality (i.e., appropriate section thickness, good post-section stain with uranyl acetate and lead citrate).
  2. Load the holder into the SEM chamber and establish good vacuum.  Navigate to the “Safe STEM” position and insert the STEM detector.  Turn on EHT (28 kV).
  3. In SmartSEM GUI, navigate to the test thin grid.  Optimize beam condition (e.g., beam and aperture alignment, objective current wobble, stigmation, focus, etc.).  Adjust magnification to ~10,000× so that the width of the field is ~25 µm.  Optimize brightness and contrast so that the histogram is centered and spread well (use histogram function in SmartSEM annotation tool or switch to ATLAS GUI to adjust B/C live).  Make note of the magnification and B/C values (these vary depending on beam condition, gain, post-section stain on the specimen, etc.).
  4. Navigate to a grid with new PEI film and optimize focus using debris or artifacts on the film surface.
  5. Adjust the magnification, brightness, and contrast to the values recorded for the test thin grid. 
  6. Acquire image of the film (frame size = 3,072 × 2,304; scan speed = 8; with line averaging n = 2).
  7. You may also want to image the film at a reduced brightness level to visualize its flaws – if these flaws (except for holes) are not visible at B/C values at which the test thin sections are imaged, then they should not affect the quality of ultrathin section images.  Thickness variation of the film can be checked by imaging it at a lower magnification (width of the field should be 100-500 µm) and lower brightness level.  If it shows many holes (several small holes are acceptable) or many streaks or tracks, then discard this batch of grids (They can be reused; see 2.1.).  See below for troubleshooting.
  8. Use the SmartSEM annotation tool to add a scale bar and to record batch number of the film and today’s date on the image.  Store the image files in an appropriate folder location.  Suggested format for the image file name = GridQC_[Film batch number].tif; e.g., GridQC_20150125-3A.tif
  9. Note the film quality on the worksheet and petri dish holding grids with newly cast PEI film.
  10. Scan the worksheet and save along with the image files.


5.  Tips and Troubleshooting

  • Select a dry, non-humid day, to coat grids. High humidity seems to diminish film quality
  • Film thickness: Film thickness is determined by the speed at which a glass slide is removed from PEI solution. Too fast a pull speed results in a thicker film (gold to purple interference color), which will not adhere to the grids very well. Too slow a speed results in a thin film (gray interference color), which will be difficult (if not impossible) to separate from the slide.  One should determine a combination of (1) speed that is comfortable and allows for single smooth pull, and (2) the concentration of PEI film that results in a medium silver interference color at this speed.   This medium silver film color is preferred for serial sections, as this thickness is rugged enough to sustain all the handling involved; yet, the resolution under the electron beam is not compromised.  Once the speed is controlled for, then note if the film continues to come out too thin or too thick. If this is the case, then the PEI solution can be made more concentrated or diluted.
  • The films do not release from the slide: (1) The slide may not be clean enough – Move onto the next slide.  (2) The film is too thin – Do not force it and move onto the next slide.  If the film thickness is in the correct range, one can try re-cutting the glass slide with razor, as described above, or try re-dipping the slide more slowly into the water.  (3) The glass slide itself is a poor releaser – Try another batch or brand of glass microscope slides.
  • Film quality:
    • Many streaks indicate that the glass slide itself, used in film coating, is casting its own impression. Try another batch or brand of glass microscope slides.
    • Holes are most likely caused by water present in your PEI solution. Make a new solution and try again on a dry day.
    • Dust particles may come from the glass slides (e.g., insufficient cleaning) or contaminants in the PEI solution.
  • No labels

Confluence Documentation | Web Privacy Policy | Web Accessibility