Light Microscopy Protocols

Optical Profilometry - Measuring pit volume from topographical projections

The principal steps are shown in this document: Measuring-pit-volume.pdf

 

Coverglass-bottom Chambers for Imaging on an Inverted Microscope

An alternative to commercially available coverglass-bottom Petri dishes and culture chambers.

 A piece of 3" x 2" plexiglass 3/8" or 10 mm thick, with the short edges slightly beveled so that the microscope slide holder can grab the plexiglass and hold it down. The piece has a rectangular hole about 2 mm  smaller than the coverslip used (such as 22x40mm coverslips). The coverslip is glued to the bottom side of the chamber with silicone caulk (clear GE Silicone II from a hardware store) - this way if the coverslip breaks, it can be scraped off with a razor blade and replaced with a new one. The coverslips are individually measured with a micrometer gauge and only those close to 0.170 mm in thickness are used (0.165 to 0.175 mm range).


The specimen, such as plant leaf tissue, is held in place with a small glass "brick" made on a glass knife maker from a leftover glass strip used to make knives for EM. One or two of those glass bricks are enough to flatten and press the leaf down against the coverslip.

glass-bottom plexiglass chamber

 

Acetolysis Procedure for Pollen Extraction

This procedure according to Erdtman (The acetolysis method. Svensk Botanisk Tidskrift, 1960 54: p. 561-564)

 destroys and extracts everything except for exine, the highly resistant outer shell of pollen that bears characteristic morphological features used in pollen identification. The extracted pollen can then be infiltarted with suitable mounting medium for light microscopy.  This technique has been used for high-resolution 3D imaging of pollen ( Vitha, S., V.M. Bryant, A. Zwa, and A. Holzenburg, 3D Confocal Imaging of Pollen. Microsc Today, 2010. 18(02): p. 26 - 28).

All processing is done in 15ml polypropylene conical tube.

  1. Add 5-10ml 5% w/v KOH, incubate at 80° C for 10 min, stirring every 2 min.
  2. Optionally, for material with debris, filter the suspension through a nylon mesh, with pore size larger then the size of pollen to be recovered.
  3. Centrifuge at 1000x g for 5 min.
  4. Discard the supernatant, resuspend the pellet in 3-5 ml concentrated HCl by vortexing or stirring, add water to 15 ml.
  5. Centrifuge at 1000x g for 5 min.
  6. Discard the supernatant, add 10ml Glacial Acetic Acid, vortex to resuspend the pellet.
  7. Centrifuge at 1000x g for 5 min.
  8. Decant the supernatant, vortex to resuspend the pellet in the remaining liquid.
  9. Slowly add 10 ml acetolysis mixture (90 % v/v acetic anhydride, 10% v/v sulfuric acid), stirring frequently.
  10. Incubate at 80° C for ~ 7 min, stirring every 2 min.
  11. Add 1-2 ml glacial acetic acid, stir and centrifuge as before.
  12. Decant, vortex to resuspend the pollen in the remaining liquid.
  13. Store

 

Poly-L-Lysine Coated Slides

Poly-L-lysine coating improves adherence of tissue sections to the glass slides. This is important for immunolocalization, where the tissue on the slide undergoes lengthy and sometimes harsh treatment and thus tends to be lost during the process.

Reagents/Supplies: Sigma P8920 Poly-L-Lysine solution (0.1% w/v) Diluted 1:10 in distilled or ultrafiltered water to make working solution, store refrigerated in a plastic container. Use for no more than about 25 slides per 50 ml of the working solution. Slides: VWR Micro slides 4830-036 (other brands and flavors may be OK, but among those we tested the above worked the best).

  • Clean slides in acetone for 5 minutes.
  • Remove slides and wipe with a kimwipe (wear gloves).
  • Air dry slides at an angle (leaning on a tube rack or other suitable support) on paper towels.
  • Dip slides in 1:10 diluted Poly-L-Lysine solution in a plastic coplin jar for 15 minutes.
  • Air-dry slides at an angle on paper towels.
  • Mark the top right corner with a diamond edged pen.
  • Bake the slides for 1 hour at 55°C.

Gelatin-Subbed Microscope Slides

To improve adherence of semi-thin resin section to glass slides for staining, use the following procedure:

  • Make 0.5 % w/v gelatin (Knox, the grocery store jello stuff) in distilled/ultrafiltered water. Dissolve the gelatine while stirring on a heated stir plate (low heat setting)
  • In the meantime, etch pre-cleaned frosted microscope slides in 5% H2SO4 for 5 min, stirring occasionally with an applicator stick and making sure the slides are not stuck together.
  • Wash well (many times) in distilled water, again stirring the slides. Keep the slides under water, do not allow to dry.
  • Fill a Coplin jar with the gelatin solution and immerse the slides for a minute or so, then remove and place in a slide rack, let dry overnight.
  • Store the slide in a storage box.

Gatenby's Glue Subbed Microscope Slides

To improve adherence of cryosections for subsequent immunostaining.

  • Prepare Gatenby’s glue: 27% v/v ethanol, 6.3% v/v acetic acid, 1.35% w/v gelatin (Knox food-grade gelatin from the grocery store is OK), 0.09% w/v chrom alum (KCr(SO4)2). Stir without heating until the gelatin dissolves (may take several hours).
  • Etch pre-cleaned frosted microscope slides in 5% H2SO4 for 5 min, stirring occasionally with an applicator stick and making sure the slides are not stuck together.
  • Wash well (many times) in distilled water, again stirring the slides. Keep the slides under water, do not allow to dry.
  • Apply a small drop of Gatenby’s glue on a slide and spread with a gloved finger. Alternatively, put a small drop of Gatenby’s glue close to one end of the slide and use the edge of another slide to drag and spread the glue across, towards the opposite end of the slide.  Let the slides dry, and store them in a storage box protected from dust. The shelf life of these subbed slides has not been established, but they should be OK for several days. 

Histochemical Staining for β-Glucuronidase (GUS) Reporter in Plants

This protocol has proven successful for many plant species and tissues. It contains several improvements over the original protocol and also deals with possible sources of errors and their elimination. Download: GUS_Localization_in_plants.pdf

Embedding Tissues in Low-melting Polyester Wax (Steedman's Wax) for Immunolabeling

Steedman's wax (Steedman, 1957) is a low melting point embedding medium for immunohistochemistry. It is used instead of the classical paraffin wax when preservation of antigenicity is important. Download a detailed protocol here: Steedman's wax.pdf 

Advantages:

  • Soluble in ethanol, no need to use hazardous solvents such as xylene.
  • Low melting point (~35 °C) allows embedding without excessive heating. Much more convenient to work with than paraffin.
  • Excellent preservation of antigenicity.
  • It is a ribboning medium, thus serial sectioning is possible.

Disadvantages:

  • Low melting point. Embedded blocks should be kept at cool or refrigerated, sectioning requires well air-conditioned rooms. Shipping the embedded blocks may be problematic in summer.
  • More expensive than paraffin (~ $40 per 1000g)
  • Section adherence to slides may be weak for certain specimens. Problematic for in situ hybridization, where most sections may be lost during the lengthy protocol.

This embedding medium has been used for immunolocalization of numerous antigens in both plant and animal tissues. As always, optimal fixation and immunostaining conditions for a particular antigen and antibody will have to be tested.

References:

  1. Vitha, S., Baluška, F., Jasik, J., Volkmann, D., and Barlow, P. Steedman's Wax for F-actin Visualization in Actin: a Dynamic Framework for Multiple Plant Cell Functions, Staiger, C.J., Baluška, F., Volkmann, D., and Barlow, P., Editors. 2000, Kluwer: Dordrecht, The Netherlands. p. 619-636.
  2. Steedman, H.F. A new ribboning embedding medium for histology. Nature, 1957. 179: p. 1345.

Microwave-assisted Fixation and Resin Embedding of Plant Roots

This protocol was developed by Ann Ellis for our clients in Plant Pathology who study parasitic nematodes. The protocol allows excellent penetration of fixatives, good structural preservation and infiltration of the specimens with the embedding resin. The embedded material can then be sectioned for either light or electron microscopy. A scientific-grade, cooled laboratory microwave (Pelco Biowave) is used throughout the fixation and dehydration to improve and accelerate the process. Nematodes, just like insects, present a challenge for fixation and embedding, because of their low permeability. This is circumvented by applying osmium-vapor prefixation. The fixation step itself includes acrolein, an often-neglected fixation agent that offers excellent penetration and fixation quality. Please note that for other specimen types this protocol may have to be modified. Download the protocol: Microwave_protocol.pdf

Outline of the protocol:

  • Osmium vapor prefixation
  • Aldehyde fixation, microwave assisted
  • Osmium post-fixation, microwave assisted
  • Washing, microwave assisted
  • Dehydration in methanol, microwave assisted
  • Resin infiltration
  • Polymerization

Digital Photomicrography with Nikon Coolpix Cameras

The relatively inexpensive Nikon Coolpix consumer digital cameras mounted on a microscope are being used successfully in many labs for routine documentation and to produce publication-quality images. The example below shows Arabidopsis leaf mesophyll cells and their chloroplasts. The image was taken using a Nikon Coolpix 4500 camera coupled with an adapter and mounted on an Olympus BH2 microscope with a 40x objective.

 

The Coolpix 995 (3-megapixel) or Coolpix 4500 (4 megapixel) are easily adapted to be mounted on a microscope via their 28mm filter thread. Both of these cameras are now discontinued, but still available on the used market. Adapters for newer cameras are also available, but the above two models are preferable because of the ease with which they are mounted to the microscope. The Coolpix 995 and 4500 cameras can be controlled from a computer via the USB or serial cables to reproducibly set the imaging parameters, namely the zoom of the camera lens (to achieve reproducible magnification). Alternatively, a remote shutter cable can be also used. Contact Stan Vitha if you are considering such setup in your lab. Check below for some relevant links.

Remote Control Driver and GUI for the Nikon Coolpix 990, 950, 880, 775, and 995 Digital Cameras, for Linux and Windows.

CoolpixPhotoMicmac. A newsgroup dedicated to the use of Coolpix cameras on a microscope. Links to software downloads.

Krinnicam. A remote shutter software for Nikon Coolpixes that run on Windows 2000/XP platforms. It features automatic image downloading through the USB connection.