Lab 6.1 Animal histology - BalzaLabNotebook - home6.1+A… · · 2011-09-06Lab 6.1: Animal histology & microtechnique ... The presentation should last approximately 30-45 minutes

Lab 6.1: Animal histology & microtechnique

BIO-331 Cell biology – Lab 6.1 Animal histology & microtechnique 1

Purpose The purpose of this lab is to introduce cytology students to basic methods of tissue processing, staining, and histology. Successful completion of this lab should produce high quality microscope slides containing sectioned and stained human tissue samples Background & introduction Histology is the study of tissues at the cellular level. Tissues are composed of cells and macromolecules secreted by the cells to form the extracellular matrix. Tissues are generally organized into four fundamental types: nervous tissue, muscle tissue, epithelial tissue, and connective tissue. Organs are generally well-organized combinations of these tissue types, with the exception of the central nervous system, which is almost completely nervous tissue. Nervous tissue histology The human central nervous system is nearly incomprehensible in complexity. It is comprised of a vast network of over 100 million neurons grouped as circuits, supported and organized by many more glial cells. As the central nervous system contains little connective tissue, it forms very soft, gel-like structures. Key anatomical structures of the central nervous system include: 1) The cerebrum, the largest and most obvious portion of the brain divided into two cerebral hemispheres. These two halves of the brain are connected by a band of fibers called the corpus callosum, which provides a communication pathway between the two sides of the brain. 2) The cerebellum is the second largest portion of the brain, which is important for coordinated movement. 3) The spinal cord provides a communicative path from the brain to the rest of the body.

Muscle tissue histology Muscle tissue is composed of cells that are specialized to generate contractile force among cells. Based on structure and function, human muscle has been categorized into three groups: 1) skeletal muscle for discontinuous voluntary contraction, 2) cardiac muscle for strong, coordinated rhythmic contraction, and 3) smooth muscle for slow and weak involuntary contraction.

Epithelial tissue histology Epithelial tissue covers and lines structures to offer protection in the case of skin, absorption in the gut, or secretion in glands. Epithelial cells are often categorized by shape. Columnar cells are tall narrow cells, while cuboidal cells are square, and squamous cells are short. Epithelial cells are usually tethered together by a series of tight junctions to limit diffusion of solutes between cells. At their basal (bottom) surface, epithelial cells are bound through hemidesmosomes to a basal lamina composed of collagen and proteoglycan. Some common types of epithelial tissue include: 1) Simple squamous epithelia a thin cellular layer that transports fluid from blood capillaries, or gas from the lung epithelia. 2) Simple cuboidal epithelia that usually lines small ducts and tubules, which may have excretory, secretory, or absorptive functions, for example the collecting ducts of the kidney. 3) Simple columnar epithelial such that performs primarily absorptive function in the gall bladder (to concentrate bile) or in the intestine to absorb fluid. 4) Pseudostratified columnar ciliated epithelial tissue offers protection and is able to mediate transport of particles trapped in mucus out of the air passages in the bronchi, trachea, and nasal cavity. 5) Stratified squamous keratinized epithelial cells that coat the epidermis to prevent desiccation.

Connective tissue



Connective tissue is unique in that cells are only a minor component. The majority of connective tissue is comprised of extracellular matrix (ECM). The ECM connects and binds tissue together to provide support and elasticity. While cells are less common in connective tissue, fibroblasts, adipose cells, mast cells, and macrophages are commonly observed. Connective tissue is found in diverse areas of the human body including 1) the fibrous connective tissue rich in collagen that is found in tendons and organ capsules. 2) the elastic connective tissue found in the dermis of skin and the bladder that confers elasticity of these tissues 3) the specialized type of connective tissue that stores fat is called adipose tissue. Adipose tissue is comprised mainly of adipose cells. Subcutaneous layers of adipose tissue acts as “shock absorbers in the soles of the hands and palms of the feet. It also “fills” space between other tissues and acts to keep organs in place. Adipose tissue commonly has a rich blood supply.

Assignment summary In this lab you will gain experience with tissue histology. The goal of this lab is to characterize human cadaveric tissue biopsy samples at the histological and cellular level. The class will work in groups of ~3 students for this project. Each group will present their results and conclusions in a PowerPoint or Keynote based lecture format. The presentation should last approximately 30-45 minutes. Individuals within the group should be responsible for preparing slides, obtaining digital micrographs at various magnifications, and presenting at least one type of their groups assigned tissue. See Appendix C for representative results. The presentation should introduce the form and function of the tissue type in addition to a brief description of the student-prepared micrographs. The presentations will be held, Thursday April 30 at 12:30 PM in room S203. Groups will be comprised as follows:

Group 1 Nervous tissue (cerebrum, cerebellum, spinal cord, nerve, and dorsal root ganglia if easily accessible)

Group 2 Muscle tissue (cardiac, smooth, and skeletal)

Group 3 Epithelial tissue (epidermis, vasculature, lung, kidney, digestive tract)

Materials and methods Fixation The cadaveric tissue has already been fixed with formaldehyde to preserve the tissue. Fixatives kill the tissue, any microbes that might cause the tissue to rot, and kill potentially infectious pathogens. Unfortunately, the formaldehyde fixative also leaves a toxic residue, so use great caution when handling fixed tissue. After you obtain a biopsy sample from your tissue, store the biopsy in 50-70% ethanol to prevent desiccation until processed for histology. Dehydration The specimen must be dehydrated to mount in paraffin wax. This must be done gradually so that delicate tissues are not distorted. Therefore tissue samples should be dehydrated gradually through a series of alcohol/water solutions of increasing strength. Small tissue samples dehydrate faster than large tissue samples. The rate of fluid exchange can be increased with up to 30% with gentle agitation. Gentle heating up to 45oC may also increase the rate of fluid exchange, but must be used cautiously, as heat may unevenly shrink or harden the tissue. Timing is important here, as excessive dehydration will lead to hard, brittle tissue and inadequate dehydration will prevent infiltration of the clearing agent and wax. A typical time schedule for the dehydration of small (<5mm in diameter) is as follows:



Dehydration schedule: 70% ethanol 5 min 80% ethanol 5 min 90% ethanol 5 min Optional: stain tissue for 15 min with 1% Eosin in 95% EtOH to make it easier to visualize during embedding and sectioning. 95% ethanol 5 min 100% ethanol 5 min 100% ethanol 10 min Note: xylene is used in subsequent steps as a “clearing agent”…it makes the tissue more translucent, and is miscible in both 100% ethanol and paraffin wax. Xylene is potent neurotoxin, avoid contact and use under the fume hood. 100% xylene 10 min 100% xylene 10 min 100% xylene 10 min Optional: performing the next few steps under vacuum will increase the rate of tissue infiltration by paraffin, as xylene is much more volatile than paraffin. This is especially helpful with porous or dense, fatty tissue. 50% xylene / 50% paraffin wax 20 min @56oC 100% paraffin wax 30 min @56oC 100% paraffin wax 30 min @56oC Tissue embedding Position the tissue in a paraffin wax mold so that when sectioned, it may be viewed transversely or longitudinally as required. Allow the wax to completely harden before sectioning. The wax hardening process may be accelerated by cooling on cold plate. This has the added benefit of producing a small wax crystalline structure, yielding fewer sectioning artifacts. The embedded tissue may be safely stored for years at 4oC until sectioning. Trimming the block Before sectioning the embedded tissue, the paraffin block may be trimmed to maximize the number of sections that may fit on a slide. To trim, cut away the excess paraffin using a safety razor blade. Use caution to not cut your fingers. Do not trim too close to the tissue (leave about 5mm of paraffin surrounding the tissue on all sides. It is important when trimming the block to leave the top and bottom sides parallel to ensure that the ribbon of sections will come off the microtome straight (if you notice the ribbon bending or curling, retrim the block). Tissue sectioning Microtome blades are exceedingly sharp and dangerous. Always exercise great care when using the microtome. Never allow your fingers to get anywhere near the blade edge. Never mount your paraffin block with the blade mounted, as you could easily slip and severely cut yourself while mounting the block. Instead, always remove the blade when mounting or adjusting the paraffin block. When sectioning, use a sharp and clean disposable microtome blade (blade can be cleaned with a q-tip dipped in xylene). The rotary microtome (Figure 1A) should be set to cut 4-12 µM (8-10



µM is a good starting thickness for novice histologists) sections, the blade clearance angle should be at approximately ~5o (3-8o). Increase the blade angle if you observe alternating thick and thin sections. Decrease the blade angle if you observe chatter (thick and thin zones in each section parallel to the blade edge). As you begin to section, first ensure that the feed mechanism is reset using the crank on the left of the microtome (optimally the crack should be about ½ inch away from all the way in). Be patient when starting a ribbon of sections, it is common to loose the first few sections. Once a ribbon starts, support the end of a ribbon with a camel’s hair paintbrush in your left hand. Use your right hand to turn the crank at a relatively slow yet steady speed. Slower speeds may result in cleaner sections. Carefully transfer the ribbon to a piece of cardboard. At this point you may cut the ribbon into lengths appropriate for mounting on microscope slides. To cut the ribbon, you may use a safety razor blade, a scalpel, or paint brushes. Use what works best for you. Do not place the ribbon on the surface of the table, as the section may stick. Use caution to avoid air currents (even deep breaths) which may disturb your sections.

Mounting sections Float the sections in a warm water bath (Figure 1b) until the ribbons expand and lay completely flat. This step is important, because sectioning frequently compresses and distorts the tissue. Bath should be set at 36-42oC (5-10o below the melting temperature of paraffin). Be sure the bath is about ¾ full. Using distilled water may help to prevent the formation of damaging water bubbles in the bath. A small amount of alcohol or a drop of detergent may reduce the surface tension, and facilitate more rapid section flattening. The sections should be fully flattened in about 1-2 minutes. Use caution to not float the sections for too long as this may cause tissue expansion and distortion. To maintain the sections in proper order while floating one may “anchor” the sections by touching them to the side of the glass water bath. When the sections appear completely expanded scoop up sections on clean pre-washed microscope slides. For optimal tissue adherence to slides use electrostatically charged “plus” slides. These slides offer superior tissue adherence. Be sure to label the slides carefully using

Figure 1. A parrafin block mounted on the microtome. Use great caution the microtome blade is very sharp and dangerous! (Photo by Karissa Pryes, 2011)



pencil, as the solvents used in the staining procedure will dissolve any ink. The slides containing sections may be stored at 4oC until you are ready to stain.

Figure 2. A) Microm HM325 rotary microtome. B) Lighted tissue floatation bath.

Hematoxylin & Eosin Stain1 (required): The most widely used stain for the visualization of animal tissue is hematoxylin and eosin (See Figure 2). Eosin stains, the positively charged cytoplasmic constituents and connective-tissue fibers light red or pink. Hematoxylin stains the negatively charged DNA and other macromolecules blue.

Figure 3. Micrograph (400x) of a human bone marrow biopsy illustrating typical hematoxylin and eosin staining (John Chol, 2004). First dry the slides on a heat block set 58oC for 10-15’ to remove any water from the sections, soften the paraffin, attach tissue to slide, and aid in the spreading out of sections to remove folds. Alternatively, the sections may be heated overnight at 37oC. Use caution to not overheat the sections as this may result in cellular distortion. The sections may be heated on an electronic heat bock with the block inverted as shown in figure 3.

1 This protocol is adopted from Bancroft & Gamble (2008) Theory and Practice of Histological Techniques, 6th Ed. Elsevier Press. p. 126-127.



Figure 4. Example of a digital heatblock used to dry the slides. Be sure to invert the metal blocks as shown on the top. 1. Deparaffinize the sections using the following series:

xylene 5 min xylene 5 min 100% ethanol 2 min 100% ethanol 1 min 95% ethanol 1 min 80% ethanol 1 min 70% ethanol 1 min water 1 min Stain with Ehrlich’s hematoxylin 20-45 min

Ehrlich’s hematoxylin solution: Dissolve hematoxylin in alcohol first, then add the other chemicals (as listed below). Allow to ripen ~2 months in a sunny windowsill before use. Filter before first use.

Hematoxylin 2g Absolute alcohol 100ml Glycerol 100ml dH2O 100ml Glacial acetic acid 10ml Aluminum Potassium Sulfate (potassium alum) ~15g

2. Wash in gently running tap water (do not use house dH2O, it’s too acidic for this application) for <5 minutes until the water runs clear and the tissue “blues.” 3. Differentiate in 1%HCl in 70% EtOH for 5-10 seconds (keep slide in gentle motion) 4. Wash again in gently running tap water for 10-15 minutes until the water runs clear and the tissue “blues” 5. Stain with 1% Eosin Y for ~10 min at RT.

EosinY solution: dilute 5g EosinY in 500ml of DI water, add a crystal of thymol to inhibit fungi, and add 250uL acetic acid to provide sharper staining)

6. Wash in gently running tap water for 1-5 minutes until the water runs clear

7. Dehydrate according to the schedule below:

70% ethanol 1 min



80% ethanol 1 min 90% ethanol 1 min 95% ethanol 1 min 100% ethanol 1 min xylene 1 min xylene ~5 min

8. Coverslip slides using Permount (use caution, this solution is xylene based). -Place a drop of Permount on the slide using a glass rod. -Angle the coverslip and let fall gently onto the slide.

-Allow the Permount to spread beneath the coverslip, covering all the tissue. -Allow to dry overnight in or near the fume hood.

Masson’s Trichrome Stain2 (optional): Specialized stains are required to visualize collagen and other connective tissues. Masson’s trichrome stain (see Figure 5) is a commonly used stain for labeling collagen. Masson’s trichrome labels nuclei black, cytoplasm red, and collagen bluish-green.

Figure 5. Micrograph (100x) of a mouse spinal cord section illustrating typical Masson’s Trichrome stain (Kayla Chastain, 2005). CAUTION: Bouin's solution (375mL saturated aqueous picric acid, 125mL formalin, and 25mL glacial acetic acid) is hazardous and the picric acid, when in less than 10% water, is very explosive. Use caution. If you observe any salt crystals on the picric acid or Bouin’s solution dissolve crystals under running water. These crystals may be explosive. First bake the slides on a heat block set 58oC for 10-15’ to remove any water from the sections, soften the paraffin, attach tissue to slide, and aid in the spreading out of sections to remove folds. 1. Deparaffinize the sections using the following series:

xylene 5 min xylene 5 min 100% ethanol 2 min 100% ethanol 1 min 95% ethanol 1 min 80% ethanol 1 min

2 This protocol is adopted from Bancroft & Gamble (2008) Theory and Practice of Histological Techniques, 6th Ed. Elsevier Press. p. 120.



70% ethanol 1 min water 1 min

2. Mordant (treat with chemicals that will assist in precipitating stain) in Bouin's Solution at room temperature overnight in a hood (do not mordant >24 hours as this will stain the tissue yellow). Spent Bouin's solution should be placed in an appropriate waste container. 3. Wash slides in gently running tap water to remove yellow color from sections. 4. Stain the nuclei blue-black in Weigert's Iron Hematoxylin Solution for 5 minutes (one may not use Ehrlich’s hematoxylin as it is sensitive to the acidic solutions used in the trichrome protocol.

Weigert’s Iron Hematoxylin solution (make fresh at least every 10 days): add equal volumes of Solution A (1% Hematoxylin in 95% EtOH) and Solution B (1.2% Ferric Chloride and 1% Acetic Acid in distilled water).

5. Wash in gently running tap water for <5 minutes until the water runs clear and the tissue “blues.” 6. Differentiate in 1%HCl in 70% EtOH for 5-10 seconds with gentle aggitation. 7. Wash again in gently running tap water for 10-15 minutes until the water runs clear. 8. Stain the cellular cytoplasm or muscle protein red in Biebrich Scarlet-Acid Fuchsin for 5 min at RT. 9. Rinse in diH2O. 10. Place the slides in Phosphomolybdic/Phosphotungstic Acid Solution for 5-10 minutes to allow for the uptake of the Aniline blue stain.

Phosphotungstic/Phosphomolybdic Acid Solution (make fresh): mix 1 volume of Phosphotungstic Acid Solution and 1 volume of Phosphomolybdic Acid Solution with 2 volumes of distilled water. Discard after one use. Don’t worry about precipitate in Phosphomolybdic Acid Solution, as this will not affect performance.

11. Stain collagen Extracellular matix with Aniline Blue Solution for 5 minutes. 12. Rinse in diH2O. 13. Differentiate in 1% acetic acid solution (make fresh) for 2-5min.

Discard this solution after use. This acid differentiation should render the shades of color more delicate and transparent. If left in the acid too long the tissue stain will appear faded and pale.

14. Dehydrate the section according to the following schedule:

70% ethanol 1 min 80% ethanol 1 min 90% ethanol 1 min 95% ethanol 1 min 100% ethanol 1 min xylene 1 min xylene ~5 min

15. Coverslip slides using Permount (xylene based). -Place a drop of Permount on the slide using a glass rod.



-Angle the coverslip and let fall gently onto the slide. -Allow the Permount to spread beneath the coverslip, covering all the tissue. -Allow to dry overnight in or near the fume hood. (If Permount is oozing out the side of the slide, don’t touch it or allow it to come into contact with a microscope stage or objective! Instead, all ow the mountant to dry overnight and then remove the excess carefully with a razor blade.

Bielschowsky’s Silver Stain3 (optional): Specialized stains are useful for staining neuronal axons, dendrites, and synaptic structures. Bielschowsky’s silver stain (see Figure 6) is commonly used for this purpose. Bielschowsky’s silver stain labels the neurofibrils black and the background brown.

Figure 6. Micrograph (100x) of a transverse section through a human peripheral nerve illustrating Bielschowsky’s silver stain (Mike Reep, 2009). First bake the slides on a heat block set 58oC for 10-15’ to remove any water from the sections, soften the paraffin, attach tissue to slide, and aid in the spreading out of sections to remove folds. 1. Deparaffinize the sections using the following series:

xylene 5 min xylene 5 min 100% ethanol 2 min 100% ethanol 1 min 95% ethanol 1 min 80% ethanol 1 min 70% ethanol 1 min water 1 min

2. Cover the sections with ~1mL Silver A for 1-2h at 37oC in a moist chamber. Silver A: 20% silver nitrate (dilute 10g in 50mL diH2O). 3. Wash slides twice in diH2O (about 1min per wash). 4. Incubate in a pot of Reducer A for 3-5min at RT with gentle aggitation. This should cause the tissue to turn yellow.

Reducer A (make fresh): 50mg pyrogallol, 5mL formalin into 600mL 80% EtOH. This

3 This protocol is adopted from Bancroft & Gamble (2008) Theory and Practice of Histological Techniques, 6th Ed. Elsevier Press. p. 369-370.



solution should be changed after every pot. 5. Wash slides three times in diH2O (about 1min per wash). 6. Place cells in Silver B solutionf for ~30 seconds (prolonged treatment may cause precipitation).

Silver B (make fresh): To 20mL of Silver A add 20mL of 95% EtOH. Add ammonia until precipitate formed is redissolved. Add 10 more drops of ammonia.

7. Briefly wash slides in diH2O. 8. Incubate in a pot of Reducer B for 2-5min at RT with gentle aggitation.

Reducer B (make fresh): 5mL 37% formalin in 95mL diH2O.

9. Briefly wash slides in diH2O. 10. Check slides microscopically. If necessary, repeat Silver B and Reducer B treatment (steps 6-9) to darken stain if necessary. 11. Tone in gold chloride for 2-3min. Gold chloride solution: (0.2% solution from RowleyBio.com # K-691-5) 12. Briefly wash slides in diH2O. 13. Fix in sodium thiosulfate solution for 5min. Sodium thiosulfate solution: to make 5% solution, dissolve 25g in 500mL diH2O. 14. Briefly wash slides in diH2O. 15. Dehydrate the section according to the following schedule:

70% ethanol 1 min 80% ethanol 1 min 90% ethanol 1 min 95% ethanol 1 min 100% ethanol 1 min xylene 1 min xylene ~5 min

16. Coverslip slides using Permount (xylene based). -Place a drop of Permount on the slide using a glass rod. -Angle the coverslip and let fall gently onto the slide.

-Allow the Permount to spread beneath the coverslip, covering all the tissue. -Allow to dry overnight in or near the fume hood. (If Permount is oozing out the side of the slide, don’t touch it or allow it to come into contact with a microscope stage or objective! Instead, all ow the mountant to dry overnight and then remove the excess carefully with a razor blade.

Movat’s Pentachrome Stain4 (optional): Movat’s pentachrome stain is complex, yet beautiful (see Figure 7). This techniques is especially useful for gut, lung, and vasculature. Movat’s pentachrome stain labels nuclei black; mucin &

4 This protocol is adopted from Luna (1968) Manual of Histolgic Staining Methods 3rd Ed. McGraw Hill Book Co. p. 95.



background blue; collagen & connective tissue yellow; muscle & fibrin red; and elastic fibers purple.

Figure 7. Movat pentachrome stain from a small intestine section (adopted from Bancroft & Gamble, p. 157). First bake the slides on a heat block set 58oC for 10-15’ to remove any water from the sections, soften the paraffin, attach tissue to slide, and aid in the spreading out of sections to remove folds. 1. Deparaffinize the sections using the following series:

xylene 5 min xylene 5 min 100% ethanol 2 min 100% ethanol 1 min 95% ethanol 1 min 80% ethanol 1 min 70% ethanol 1 min water 1 min

2. Stain with mucin and background substace with alcian blue for 20min. Alcian blue solution: to make 1% solution, dissolve 5g Alcian Blue (8 GS) in 490mL diH2O and 10mL glacial acetic acid. 3. Wash in gently running tap water for ~5 minutes until the water runs clear. 4. Place slides in alkaline alcohol for 1-2h (this forms the insoluble pigment monastral fast blue from the alcian blue). Alkaline alcohol solution: 50mL ammonium hydroxide into 450mL 95% EtOH. 5. Wash in gently running tap water for at least 10 minutes.

NOTE: this step is important as incomplete removal of the alkaline alcohol will inhibit subsequent steps.

6. Briefly rinse slides in 70% EtOH. 7. Stain the elastic fibers purple in Resorcin fuchsin solution for 12-24h.

Resorcin fuchsin solution (make fresh): add 10mL of Resorcin fuchsin stock (RowleyBio F-384-3) to 100mL of 70% EtOH and 2mL concentrated HCl.

8. Briefly rinse slides in diH2O. 9. Stain the nuclei and elastic fibers black in Weigert's Iron Hematoxylin Solution for ~15min (do not use Ehrlich’s hematoxylin as it is sensitive to the acidic solutions used in the Movat



protocol). Weigert’s Iron Hematoxylin solution (make fresh at least every 10 days): add equal volumes of Solution A (1% Hematoxylin in 95% EtOH) and Solution B (1.2% Ferric Chloride and 1% Acetic Acid in distilled water).

10. Wash in gently running tap water for ~5 minutes until the water runs clear. 11. Briefly rinse slides in diH2O. 12. Stain the muscle and fibrin red in Woodstain scarlet-acid fuschin solution (RowleyBio F-384-4) for 5 min. 13. Briefly rinse slides in 0.5% acetic acid solution.

0.5% acetic acid solution: add 2.5mL concentrated acetic acid to 500mL diH2O. 14. Differentiate in 5% phosphotungstic acid solution for 10-20 minutes. Thes background should be blue and the collage pale pink. 5% phosphotungstic acid solution: dissolve 25g phosphotungstic acid in 500mL diH2O. 15. Briefly rinse slides in 0.5% acetic acid solution.

0.5% acetic acid solution: add 2.5mL concentrated acetic acid to 500mL diH2O. 16. Rinse the slide three times with 100% EtOH (it is important to remove all traces of water before staining with the very hydrophobic safran, so use the high purity EtOH from Sigma, not the cheap histological grade EtOH). 17. Stain the collagen and connective tissue yellow in alcoholic safran solution (RowleyBio F-384-5) be sure to keeo the cap on the alcoholic safran stock solution tight. 18. Rinse the slide three times with 100% EtOH to remove excess safran. 19. Rinse the slide two times in clean xylene (use caution). 20. Coverslip slides using Permount (use caution as Permount is xylene based). -Place a drop of Permount on the slide using a glass rod. -Angle the coverslip and let fall gently onto the slide.

-Allow the Permount to spread beneath the coverslip, covering all the tissue. -Allow to dry overnight in or near the fume hood. (If Permount is oozing out the side of the slide, don’t touch it or allow it to come into contact with a microscope stage or objective! Instead, all ow the mountant to dry overnight and then remove the excess carefully with a razor blade.



Appendix A: Trouble shooting guide Histology microtechnique is as much an art as it is a science. Many of these skills are difficult to describe in written protocols. They may only be acquired through practice. Simple solutions to common problems with microtechnique are shown below. Be patient. Don’t get discouraged too quickly. Persistence in histology is often rewarded with incredible results. Problem Cause (solution) Ribbon is curved Edges of block are not parallel (re-trim block) Alternating thick and thin sections Paraffin too soft for tissue (cool paraffin)

Insufficient blade angle (increase clearance angle) Faulty microtome mechanisms (tighten block & blade)

Alternating thick and thin zones parallel to blade edge in each section (chatter)

Loose blade or block (tighten) Excessive blade angle (decrease clearance angle) Over-dehydration or hardening (soften with warm lamp or warm breath) Dull blade (replace) Calcified tissue (decalcify or use less calcified tissue)

Tissue splitting at right angles to the blade Nicks in blade (use different part or replace blade and avoid dulling the blade with hair from the paint brush in the future) Visible mineral deposition (remove with a fine forceps or scalpel)

Sections will not form ribbons Paraffin too hard (soften with warm lamp or warm breath)

Sections compress too much, distorting tissue or folding

Paraffin too soft (harden by cooling on ice)

Sections roll into a tight coil making it difficult to start a ribbon

Sections are too thick (reduce section thickness) Excessive blade angle (decrease clearance angle)

Sections attach to block on return stroke Static electricity (humidify air or decharge with dryer sheets) Insufficient blade angle (increase clearance angle) Debris on blade or block (inspect, clean and/or replace)

Incomplete sections or sections with holes Incomplete impregnation of the tissue with paraffin (melt and reprocess) Tissue may be falling off due to non-electrostatically charged slides (use positively charged slides)

Sections disintegrate on water surface Water float bath too hot (reduce temperature – this may be done quickly by dropping in a couple ice cubes)



Appendix B: Oral presentation assessment rubric Criteria Level of performance

A = superior & sophisticated

B = good C = average & unremarkable

D = in need of improvement

F = failure / unacceptable

Introduction (10pts)

Fascinating summary of the state of the field, a clear articulation of goals, and evidence of comprehensive research

Generally clear and interesting but contains minor ambiguities or omissions

Competent but boring (failed to capture the attention of the audience), some evidence of inadequate research

An explanation of the importance and background of the study is attempted, but inaccuracies and omissions are obvious

Background information is absent or misrepresented, importance of the study is assumed, goals are unclear

Results & methods (10 pts)

Structured and lucid articulation of the methods and outcome

Generally coherent, but difficult to understand at times

While logically arranged, poor transitions result in unnecessary confusion

Obviously disorganized, but remains somewhat comprehensible

The level of disorganization renders the results incomprehensible

Conclusion (10 pts)

Logically consistent and flows seamlessly from the presented results

Generally consistent with some minor ambiguities

Vague or weak conclusion leaves the audience wanting, ends abruptly

Contains inconsistencies or over-reaching claims

Obviously inconsistent with presented results

Style & delivery (10 pts)

Delivered with charisma, dynamic, and even exciting at times. The research “comes alive” and ends on time

Delivered with confidence, good eye-contact with the audience, but lacks some measure of polish, may finish a few minutes late

Generally articulate, but obviously nervous, merely “reads” notes or slides, eye-contact with the audience appears forced

Fails to engage the audience, long uncomfortable pauses, and frequently caught searching for the “right” word. Either wastes the allotted time or far exceeds the allotted time

Unprofessional, frequently mispronounced words, total dependence on notes or slides, sloppy appearance detracts from credibility

Organization (10 pts)

Evenly distributed workload, well organized throughout. A cohesive presentation

Evenly distributed workload, generally well organized, only occasionally losing focus

Generally organized, but some aspects of the presentation are disjointed

The presentation barely hung together, many aspects of the presentation seem only slightly related to one another

Various topics appear “out of nowhere,” making it almost impossible to follow the presentation

Visual media (10 pts)

Professional appearance and masterfully described visuals with more images than words.

Professional appearance and competently described, some slides may appear too “wordy”

Of some value, but appear overcrowded or haphazard in construction, some slides contain dense text

Distracts and confuses rather than informs, some slides are illegible

Necessary or useful visual media is absent or failed for technical reasons

Histology (40 pts)

Appropriate embedding orientation, artifact-free sections, balanced stains, and professional micrographs

Appropriate orientation, artifact-free sections, but stains not fully optimized, micrographs slightly unfocused

Sections may contain minor artifacts, suboptimized stain and unprofessional micrographs…but histology retains considerable value

Confusing orientation, may contain major artifacts. Only one stain attempted. Poor micrograph quality

Failed to demonstrate abiity to obtain micrgraphs of sectioned and stained tissue



Appendix C: Representative student results

Figure 15 Transverse 10µm sections of formalin-fixed and paraffin-embedded human nerve tissue. (A) Micrograph obtained using the 20x objective to visualize hematoxylin and eosin stained tissue. (B) Same as (A) but visualized using the oil-immersion 100x objective. (C) Same as (A) but silver stained. (D) Same as (C) but visualized using the oil-immersion 100x objective.

5 These micrographs are of human tissue sectioned and stained by Mike Reep in the Spring 2009 course.

Documents

Lab 6.1 Animal histology - BalzaLabNotebook - home6.1+A… · · 2011-09-06Lab 6.1: Animal histology & microtechnique ... The presentation should last approximately 30-45 minutes