Histological Techniques

Transcription

1 Histological Techniques Histology is the study of the cellular organization of body tissues and organs. The term is derived from the Greek "histos" meaning web or tissue, and refers to the "science of tissues". Reference to anatomical structures as "tissues" originated with the French surgeon Bichat, who compared the characteristic appearance of different parts of the body to the texture of cloth ("tissue derives from the Latin word "texere", to weave). The light microscope is the tool used most widely for clinical applications of histology. However, the advent of the electron microscope greatly extended the detail at which subcellular structure can be studied. Thus, histology now embraces the study of the structures of both tissue and cells, and the relationship between these structures and physiological function. The structure of cells and tissues can be distinguished at two levels. The fine structure is that which can be distinguished at the level of light microscopy (a magnification of 1000 x or less). Electron microscopes are generally employed to study ultrastructure B the detailed structure of the cell cytoplasm, organelles and membranes that is not discernable with a light microscope. Many techniques have been developed which are designed to preserve the structural integrity of a specimen so that it can be viewed microscopically. The process through which cell structure is preserved is called fixation. Since cells rapidly deteriorate after a tissue has been removed from the body, achieving adequate fixation is often the most difficult task confronting a histologist. "Artifacts" are changes to the original structure of cells and tissues that arise from tissue deterioration and from the fixation process itself. Thus, a skilled histologist employs techniques that minimize the formation of artifacts in different types of tissues, and has is the ability to distinguish artifacts from normal cell structures. Cell structure is most commonly studied in slices of the tissue, called sections, that are thin enough to allow transmission of light or an electron beam. There are many methods of sectioning tissues, and sometimes particular tissues require special techniques. The method most widely employed is called the paraffin method. Although this technique is not universally applicable, e.g. it does not work well with hard tissues such as woody parts of plants or bones from animals, it does present many advantages over alternative methods. The necessary reagents are inexpensive, readily available, and much less toxic to humans than those used in most other techniques. This laboratory exercise will be performed over the period of three weeks. Working in groups of two, you will prepare sections of mouse tissue and produce photomicrographs and descriptions of your observations. It will be necessary for you to return during periods outside of the regular laboratory periods to complete this exercise. This is unavoidable, but the timing of many of the steps in the procedure is flexible enough that you and your partners will be able to fit these procedures into your schedules. OBJECTIVES Histology Page 1

2 1. To learn and understand the steps of the paraffin method of section preparation. 2. To prepare thin sections of an organ and learn something about its cellular structure. 3. To understand the importance of distinguishing "artifacts" from meaningful cell structure. 4. To demonstrate an ability to work carefully and meticulously in the lab 5. To learn how to prepare publication quality images. We will be sacrificing a mouse in this lab exercise as a source of tissue. Because no laboratory animal should be sacrificed without suitable justification, you will be expected to direct your utmost attention to achieving the educational goals of this laboratory exercise. THE PARAFFIN METHOD All histological procedures can be divided into a similar series of steps. For the paraffin method these steps are as follows: I. Tissue resection II. Fixation III. Dehydration IV. Infiltration and embedding in paraffin V. Sectioning with a microtome VI. Mounting on microscope slides VII. Clearing and Staining VIII. Preparation of permanent mounts You will be expected to complete steps I - IV during the first week. Details about these steps and the procedures you will follow are given below. Histology Page 2

3 I & II. Tissue resection and fixation. We will sacrifice (kill) mice to remove (resect) certain organs the lungs, kidneys and liver. The lungs and kidney will be used to learn the steps of paraffin sectioning, the most widely used method for doing histological analysis. The kidneys will be used as a source of protein to be analyzed by gel electrophoresis and immunoblotting. There are several ways in which a small rodent can be humanely killed for use in biological research, each with particular advantages and disadvantages. Mice may be euthanized by inhalation of carbon dioxide, methoxyflurane, or halothane. If exposure to chemicals contradicts the objectives of the investigation, cervical dislocation (rupturing of the spinal column in the neck) can be performed. In this lab the instructor will use a rodent guillotine to decapitate sedated mice. This is instantaneous and painless, and allows bleeding out of the blood, which otherwise fills the body cavity during dissection. It also allows the organs to drain of blood that might interfere with an analysis of organ-specific proteins. Any student who has objections to working with sacrificed mice is excused from doing so personally, but should watch the procedure as performed by their partner. Supplies needed (per group of two): [Manufacturer s name is included parenthetically] Mouse Plastic sample jar (for fixative solution) Ice buck packed full of ice Plastic cryostorage vial (for the liver section) 2 Tissue cassettes ½ (top or bottom) plastic petri plate Histochoice-MB (Ameresco) - fixative Dissecting tray and surgical tools (scalpels, forceps, etc) Resection of lungs and kidneys These will be placed in fixative as the first stage of the Histology lab exercise that will be completed later in the semester. The primary function of a fixative is to preserve the cellular structure of the tissue. Fixation is necessary to protect and harden the tissue against the deleterious effects of later procedures which otherwise would disrupt cellular structure beyond recognition. Furthermore, fixation minimizes a process called autolysis. Autolysis is the degradation of cellular structure which results from the release of degradative enzymes from the excised tissue itself. The fixation process must be started as quickly as possible after removal of the sample. 1. On labeling tape, label the plastic sample jar with the names of the group members, date, and lungs and kidneys. Label the tissue cassettes in pencil as lungs and the other as kidneys 2. Fill the vial about 2/3 full with the fixative. 3. Remove the organs from your mouse. 4. Place the lungs and kidneys into separate tissue cassettes and then into the vial containing 75 ml of fixative labeled with your group=s initials and tissue type. 5. Store the vial on the bench top at your work area. Histology Page 3

4 Liver resection 1. Label a cryostorage vial on labeling tape with your names, date, and 0.5g liver 2. Resect the liver and place it on a Petri plate chilled in your ice bucket. 3. Tara a Petri plate on a scale and trim liver to obtain an approximately 0.5g piece. 4. Transfer the liver piece in the cryostorage vial. 5. Store the vial on ice until transferring it to the -80 O C freezer. III. Dehydration. After fixation, the water must be removed from the tissue block, a process called dehydration. Isopropyl alcohol (IPA) is a favored reagent because it is miscible in paraffin. The tissue must not be dehydrated rapidly because this will cause distortion of the tissue. Rather, dehydration is carried out in a slow, step-wise manner by passing the tissue block through a series of solutions of increasing IPA concentration. In this way the water is fully leached out and replaced with IPA. Supplies needed (per group of two): 70%, 85%, 95% and 100% IPA solutions that you prepare 1. Pass your tissue capsules through the following series of solutions. 2. For each step, place approximately 50 ml a fresh IPA solution into your plastic vial. 3. Decant the used solutions into the organic solvent disposal container in the hood. Step 1. 70% IPA 1 hr 2. 70% IPA 1 hr 3. 85% IPA 1 hr 4. 95% IPA 1 hr % IPA 1 hr % IPA 1 hr incubation period Equilibration Equilibration means to allow a solution to reach a stable concentration within a tissue. Thus, for example, after 1 hour the IPA will have reached 70% within the tissue block. IV. Infiltration and embedding. Prior to sectioning, the tissue block must be infiltrated with a material that acts as a support during the sectioning process. For the method described here, paraffin serves this purpose. We will be using Paraplast Plus (Fisher Scientific). During infiltration, the paraffin will equilibrate within the tissue block, eventually occupying all of the space in the tissue that originally held by IPA. After infiltration, the tissue is allowed to solidify in a mold, embedded within a small cube of paraffin. Supplies needed: Melted paraffin in metal pitchers 4 base molds Petri plates Infiltration 1. Discard the 100% IPA from the last dehydration step, and fill the vial about : full with melted paraffin. 2. Allow tissue to equilibrate for 1 hour in an incubator set at 58 O C. Histology Page 4

5 3. Pour the paraffin into the container labeled for paraffin disposal. 4. Repeat step 1 using fresh melted paraffin. Embedding 1. Place base-pieces for two embedding molds in a plastic Petri plate label the plate along the edge with your name. 2. Decant the paraffin from the second infiltration step into the waste container. 3. Working quickly but carefully, use forceps to transfer the tissue blocks to the well of separate base mold, snap the base of tissue cassette into the base mold and then fill the mold with paraffin. 4. Allow the paraffin to solidify at room temperature. *** If the paraffin begins to solidify homogeneously around the tissue block, allow the paraffin in the base mold to melt in the incubator, and then allow it to solidify. V. Sectioning. Sectioning is accomplished by using a cutting apparatus called a microtome. The microtome will drive a knife across the surface of the paraffin cube and produce a series of thin sections of very precise thickness. The objective is to produce a continuous "ribbon" of sections adhering to one another by their leading and trailing edges. The thickness of the sections can be preset, and a thickness between 5-10 μm is optimal for viewing with a light microscope. The sections can then be mounted on individual microscope slides. Preparation and mounting of the embedded tissue block on the microtome is very important to successful sectioning. The paraffin surrounding the tissue block must be first trimmed, and then secured to a holder which is then mounted on the microtome. The procedures for trimming and mounting your paraffin block, and using the microtome will be demonstrated during the laboratory period. Troubleshooting suggestions for diagnosing difficulties during sectioning are provided at the end of this exercise. VI. Mounting of sections on microscope slides. In this procedure, the sections are permanently attached to microscope slides. If "serial" sections are desired, (i.e., sections that reveal sequential layers of the tissue structure) then sectioning must be performed carefully and systematically. Supplies needed (per group of two): 10 microscope slides. Coplin jar Slide storage box Hematoxylin (Surgipath/Leica) Eosin (Surgipath/Leica) Preparing the microscope slides 1. Using a diamond pen, label the 5 microscope slides at one end with your name and tissue type, and numbered sequentially from 1 5; five slides will be used for the lung and 5 for the kidney. 2. Wash the microscope slides with soap and water, and rinse free of soap with tap water. 3. Place the slides in a coplin jar and rinse several times with roh 2 O. 4. Handling the slides only by their edges, place the slides in your slide storage box, and allow them to dry. Histology Page 5

6 Mounting sections on microscope slides It will be apparent during sectioning that the sections are not perfectly flat, but rather slightly crinkled. This is normal, and the sections will become flattened by floating them on water held at 45 O C. The solution also contains an adhesive, Sta-On (Surgipath/Leica), which causes the tissue section to bind to the slide. 1. Carefully transfer the sections to a solution held in a 45 O C water bath. Within a few seconds you should see the sections flatten and the wrinkles disappear. 2. Dip a clean microscope slide into the adhesive solution, and slowly pull it upward, out of the solution, allowing sections to adhere to the surface. Make sure that the slide is oriented with the label facing upward. 3. Dry the bottom of the slide and carefully blot excess adhesive from around the sections (be careful not to touch the sections themselves). 4. Allow the slides to dry overnight in the storage box. VII. Clearing and staining. Before a section can be stained the paraffin must be removed, a process called clearing. After clearing, only the tissue remains adhering to the slide. Clearing is accomplished by passing the mounted sections through the solvent Clearene (Surgipath/Leica) that dissolves the paraffin. Staining of histological sections allows observation of features otherwise not distinguishable. For routine histological work, it is customary to use two dyes, one that stains certain components a bright color and the other, called the counterstain, that stains other cellular structures a contrasting color. While literally hundreds of staining techniques have been developed, the two stains most widely used for routine work are hematoxylin and eosin Y (commonly abbreviated as AH & E@). (For your lab report, report the specific types of these stains that you use.) Hematoxylin stains negatively charged structures, such as DNA, a blue color. Eosin imparts a red color to most of the other cell components. To produce permanent staining with hematoxylin, the dye must be oxidized to "hematein", which is achieved by treating the tissue sections with Scott's solution. Supplies needed (per group of two): coplin jars IPA solutions Hematoxylin Eosin Clearene (Surgipath/Leica) clearing agent Scott s solution 240 mm NaHCO mm MgSO 4 Histology Page 6

7 Clear and stain your slides with the following schedule of solutions held in Coplin jars: Clearing and Rehydration 1. Clearing agent #1 3 minutes 2. Clearing agent #2 2 minutes 3. Clearing agent #3 1 minute % IPA 30 seconds 5. 85% IPA 30 seconds 6. 70% IPA 30 seconds 7. Tap water 30 seconds Staining 8. Hematoxylin 2 minutes 9. Tap water 30 seconds 10. Scott s solution 1 minute 11. Tap water 30 seconds 12. Buffer 1 minute 13. Tap water 30 seconds % IPA 1 minute % IPA 1 minute 16. Eosin Y 1 minute Rinsing, Rehydration & Mounting Prep % IPA #2 2-3 minutes % IPA #3 2-3 minutes 19. Clearing Agent 1 minute 20. Clearing Agent 1 minute 21. Clearing Agent 1 minute Each group will have their own coplin jars with the solutions high-lighted in gray. Each group should fill the Tap water coplin jars, and empty them when done. The second set of IPA and clearing agent solutions must be different than those used above. Do not allow slides to dry before mounting under cover slides. VIII. Preparing permanently mounted sections. The final step in this procedure is to permanently mount the sections under a coverslip. This is accomplished by covering the section in a medium that will harden and produce a clear binder between the slide and cover slip. The ideal mounting medium should not distort the stain color, or yellow and become brittle with age. We will use a mounting resin called Permount (Fisher Scientific).. 1. Place 2-3 drops of resin over the section. 2. To avoid entrapping air bubbles, lower the cover slip slowly from one side of the droplet. 3. Place the slide on the slide warmer and carefully place a lead weight on top of the cover slip. There should be enough mounting medium to completely cover the bottom of the cover slide, and budge slightly around the edges. 4. Leave slides on the warmer for at least 24 hours; excess medium can then be cut from edges of cover slip with a razor blade. Histology Page 7

8 IX. Photography and presentation of sections. You will photograph representative samples of your stained sections, and prepare publication quality figures. This will require that you familiarize yourself with the anatomy of your organ. A variety of histology guides and books are available for this purpose, and you are expected to use them. Each student will prepare a figure with an image of the tissue photographed under the 20x or 40x objective. The image will be annotated using Adobe Photoshop software to show: At least 4 labeled structures or cell types labeled with arrows and letters An appropriate scale bar A legend that includes a Figure number, title and suitable legend. Synopsis of steps to follow when formatting images in Adobe PhotoShop After an image is Acquired into Photoshop from the Spot Camera window, the following basic steps should be followed. Doing some of these steps in the order given will be important. This is not intended to be a thorough guide to the use of PhotoShop, some trial and error is expected on your part. 1. Save the image. Save it in your directory on the Scratch/309 directory. Save the image with a descriptive name including the objective lens magnification (e.g., Lung-1-40x ). 2. Add the appropriate scale bar. Scale bars are in the scale bars directory. Be sure to use a scale bar for the microscope (Nikon or Olympus) and objective lens that you used to take the picture. Open the scale bar file; on the Select menu select All use the copy command to copy the scale bar to the clipboard Now click on the histology image to activate that window; use the paste command to paste the scale bar into this image\ The scale bar will automatically be added on a new layer, allowing it to be repositioned as needed. You can close the scale bar window. 3. Judiciously adjust the image quality. Under Filter menu, select sharpen image, once or twice Under the Image menu, select Adjust and Brightness/Contrast to adjust these. 4. Use Text tool to add identifying letters. Text is automatically added to a new layer. 5. Use the Line tool to add arrows. 6. Save the image in a JPEG format. 7. Insert the image into a MS Word document and add suitable figure legend. Histology Page 8

9 TROUBLESHOOTING GUIDE A. Problems arising during sectioning of tissue block. I. No ribbon forms 1. Paraffin is rather "crumbly": Tissue may be incompletely infiltrated and/or contaminated with IPA or water. -- Tissue block must be re-infiltrated and embedded. 2. Tissue block may be incorrectly trimmed. -- Retrim block and make sure that all edges are at 90E angles. 3. Knife angle may be incorrect. -- Ask the instructor for help in repositioning the knife blade. II. Ribbon curves as it forms 1. Edges of block are not parallel. -- Retrim block and make sure that all edges are at 90E angles. 2. Knife blade is not uniformly sharp. -- Use a different part of knife or ask the instructor for a new knife. III. Sections are severely compressed. 1. Knife is dull. -- Try a different part of knife or ask instructor for a new knife. 2. Tissue is incompletely infiltrated or embedded. -- Tissue must be reinfiltrated and embedded. IV. Sections bulge at middle. 1. Paraffin is not of uniform temperature. -- Allow the block to equilibrate to room temperature longer. 2. Knife blade is not uniformly sharp. -- Use a different part of knife or ask the instructor for a new knife. 3. Tissue is incompletely infiltrated or embedded. -- Tissue must be reinfiltrated and embedded. V. Sections not of uniform thickness. Histology Page 9

10 1. Paraffin is not of uniform temperature. -- Allow the block to equilibrate to room temperature longer. 2. Block, or paraffin holding block to the holder, is cracked. -- Carefully examine block and holder. Remount to holder if necessary. If block is cracked, it must be re-embedded. 3. Knife is loose. -- Sure knife in position. 4. Microtome is faulty. -- Unlikely, so seek assistance from the instructor. VI. Ribbon adheres to block instead of knife. 1. Ribbon is electrified. -- See if the ribbon sticks to anything else. If so turn on a Bunsen burner or boil some water a short distance from microtome. 2. Knife angle incorrect. -- Ask the instructor for help in repositioning the knife blade. 3. Knife has paraffin pieces stuck to it. -- clean knife with xylene. 4. Block has paraffin pieces stuck to it. -- carefully scrape the block clean with a razor blade. VII. Tissue breaks away from paraffin or is shattered by knife. 1. Tissue incompletely infiltrated. -- Reinfiltrate and embed tissue. 2. Tissue is too hard for paraffin sectioning. -- Re-embed in nitrocellulose or other embedding medium. 3. Fixation or dehydration procedure inadequate for tissue. -- Different procedures would need to be used. VIII. Ribbon splits. 1. Nick in knife blade. -- Use a different part of the knife. 2. Grit in object. -- Examine edge of section carefully; If a piece of grit is apparent, then try to dissect it out or retrim the block. B. Problems arising in sections after staining and mounting. Histology Page 10

11 I. Sections distorted. 1. Dull knife. -- Tissue must be re-sectioned with sharp knife. 2. Tissues inadequately fixed, dehydrated or infiltrated. -- A more suitable procedure must be employed. II. Sections appear opaque or have highly refractive lines outlining cells and tissues. 1. Paraffin incompletely cleared from the sections. -- More thoroughly clear paraffin in subsequent sections. III. Sections will not stain, or stain irregularly. 1. Paraffin incompletely cleared from the sections. -- More thoroughly clear paraffin in subsequent sections. 2. Sections not uniform in thickness. -- See "V" above. 3. Tissues inadequately fixed, dehydrated or infiltrated. -- A more suitable procedure must be employed. Histology Page 11

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