Optimal Methods for Preparation and Immunostaining of Iris, Ciliary Body, and Choroidal Wholemounts

Abstract

purpose. Investigations into the biology of resident and infiltrating immune cells in the uveal tract of the rodent eye have been greatly aided by the use of tissue wholemount methods. These methods offer a number of advantages over conventional histological and frozen section techniques. The purpose of this article is to provide a detailed step by step guide to aid others who may wish to use this method. methods. A detailed description of whole-body perfusion fixation, dissection and isolation of the iris-ciliary body from the anterior segment and the choroid from the posterior segment is provided. In addition, the techniques used to handle whole tissue pieces during single and double immunohistochemical staining protocols, as well as the staining protocols themselves, are described. results. In refining the techniques described, the author has catalogued a number of frequent problems which compromise immunohistochemical staining results. A troubleshooting guide aimed to help identify the cause of common problems and with some suggested remedies is provided. conclusions. Although tissue wholemounts are frequently used in retinal research, a similar approach to investigating the components of the uveal tract has only recently been applied. The methods described in this article will provide sufficient detail for other investigators to obtain maximum benefit from this alternative approach and provide an additional technique to assist in their investigations of ocular immunobiology.

Investigations in the past few years in the author’s laboratory have focused on the biology of various types of immune cells in the normal and inflamed rodent uveal tract. ^{1 2 3 4 5 6} These studies have been greatly advanced by the use of tissue wholemounts combined with immunohistochemical or histochemical methods. These techniques are routinely used in preference to conventional methods, such as wax histology or frozen tissue sections. The advantages of this approach include economy of tissue preparation time, visualization of the morphology of entire cells and cell networks, and the increased sampling of tissue. The value of this technical approach relies heavily on adequate preservation of cell morphology, optimal antigen preservation, low background staining, minimization of tissue damage during dissection and handling, and careful mounting of the specimens for photography and examination. Uveal tract wholemount preparations are most informative and rewarding when applied to albino animals in which uveal tissues are sufficiently thin and transparent to allow transillumination and resolution of individual stained cells. Thus, albino rats and mice are the most commonly chosen experimental animals. Pigmented animal strains can be studied but require bleaching to allow conventional microscopic examination. Alternatively, immunofluorescence methods can be used, although problems of autofluorescence must be borne in mind. 

The purpose of this article is to provide a detailed methodologic account to aid investigators who may be considering using similar approaches. It covers all aspects of tissue preparation, from whole body perfusion, microdissection, and tissue handling, to a step-by-step guide to immunohistochemical staining. In addition, there is a supporting Internet Web site with video sequences of key procedures. 

Methods

Whole-Body Perfusion

Intracardiac perfusion can be performed by using a constant-flow pump or by gravity flow. The latter method is used in the author’s laboratory. All procedures conform to the ARVO Statement for the Use of Animals in Ophthalmic and Vision Research. 

Animals can be deeply anesthetized using a variety of methods. We use an intraperitoneal injection of pentobarbitone sodium (100 mg/kg body weight; Rhone Merieux Australia, Queensland) diluted in cold phosphate-buffered saline (PBS). When the animal is adequately anesthetized (fails to respond to a foot-pinch test) it is pinned by the feet supine to a corkboard situated inside a laminar flow cabinet. The fur on the chest and abdomen is dampened with 70% ethanol to prevent hair from contaminating wet tissue specimens. An abdominal incision is made in the midsagittal plane from the symphysis pubis to the xiphoid process. The cut is continued rostrally as paired parasagittal incisions in the chest wall, thus avoiding the paired internal thoracic arteries that lie close to the sternum. The sternum is then elevated and pinned rostrally to allow visualization of the chest cavity. Thymic tissue is displaced to expose the ascending aorta. 

The apex of the heart is grasped gently with artery forceps, and a 2-mm partial-thickness scalpel incision is made in the left ventricle. A blunted wide-bore cannula (2–3-mm internal diameter) is inserted through the incision and gently pushed rostrally until the tip is seen within the lumen of the ascending aorta. The cannula is connected by wide-bore clear plastic tubing to a three-way valve. Reservoirs of cold heparinized PBS (1 IU heparin per milliliter PBS) and cold fresh 2% or 4% paraformaldehyde are connected to the other ends of the three-way valve. These reservoirs can be elevated to varying heights by using a pulley mechanism. Elevation to approximately 1200 mm above the height of the animal ensures adequate perfusion pressure. After cannulation of the heart, the valve to the PBS reservoir is opened, and a 1- to 2-mm cut is immediately made in the wall of the right atrium to allow blood and perfusate to escape. The volume of perfusate required per animal is approximately 60 to 80 ml for mice and 250 to 350 ml for rats. After the initial PBS perfusion (approximately 5–10 minutes), virtually no trace of blood should be evident leaving the heart. The three-way valve is changed to allow fixative to enter the heart. Within 5 to 10 seconds of changing to fixative, evidence of muscular spasms should be apparent. A volume of fixative equal to that recommended for PBS should ensure satisfactory fixative perfusion. Signs of adequate perfusion include movements in the limbs and tail (indicating fixative has reached the extremities), clearing of all mesenteric vessels, blanching of the liver, and absence of pink tones in the tongue and eyes. Signs that the perfusion pressure or rate is too high include fixative flowing from the nostrils and expansion of subcutaneous tissues in the head and neck due to fluid accumulation. The animal should be rigid if the perfusion fixation has been successful. 

After completion of the perfusion, the cannula is removed from the heart, and the eyes are enucleated, by using blunt curved enucleation scissors. Other tissues may also be dissected at this time. Useful control tissues suitable for preparation as wholemounts include the non–fat-bearing region of the mesentery of the small intestine, epidermal sheets, meninges, or other thin tissues that are suitable to be flatmounted and transilluminated for examination by light microscopy. 

Microdissection of Globes and Removal of the Iris, Ciliary Body, and Choroid

Short video sequences illustrating the following steps are available (http://iaaf.anhb.uwa.edu.au/webdev/Paul/default.htm). 

Dissection of eyes is best performed under a stereoscopic dissecting microscope with a fiber-optic light source directed obliquely. This serves to reduce glare and reflections from metal dissection instruments. Dissection of albino tissues is best performed on a dark, compliant cutting surface (Fig. 1.1) . Extensive experimentation has resulted in the choice of heavy-duty, black, hard rubber blocks (approximately 10 × 6 cm and 1 cm in thickness). The globe is carefully incised (1-mm cut) immediately behind the limbus, by using a sharp razor blade (Fig. 1.2) . Curved iris scissors are then used to continue the incision around the globe and separate the eye into anterior and posterior segments (Fig. 1.3) . Jeweller’s forceps are used to hold the eyecup carefully while the lens is gently teased from the anterior segment (Fig. 1.4) . Three or four radial incisions are made with the razor blade through the anterior segment to produce pie-shaped wedges consisting of cornea–sclera externally and iris–ciliary body or choroid–retina internally (Fig. 1.5) . Excess sclera, choroid, and retina posterior to the ciliary body are carefully removed with the razor blade, using downward cutting motions (not sawing actions). Gripping the central cornea with fine forceps, a beaver blade (Tooke’s knife) is carefully placed between the inner surface of the cornea and the iris. The blade is gently pushed posteriorly into the iridocorneal angle, thus breaking any remaining adhesions between the iris–ciliary body and cornea–sclera. The isolated pieces of iris–ciliary body (Fig. 1.6) are gently lifted with either a fine artist’s paintbrush or the beaver blade and transferred into clean glass vials containing fresh PBS. 

To remove the choroid, the posterior segment (Fig. 1.7) is cut into either quadrants or thirds. The beaver blade is gently slipped under the neural retina into the subretinal space. The neural retina can then be easily removed from the underlying retinal pigment epithelium (RPE), choroid, and sclera (Fig. 1.8) . While the sclera is firmly grasped in one corner of the segment, the beaver blade is gently pushed under the choroid, which should be visible on the inner surface of the sclera. Gentle sideward movements of the blade while pushing away from the edge held by the forceps should break any adhesions between the choroid and sclera (Fig. 1.9) . The isolated segment of choroid is then lifted gently and transferred into fresh clean PBS. 

It should be noted that all dissections are performed in a fluid environment (PBS or fixative). At no stage should the tissues be allowed to dry, because it causes them to shrivel and collapse, which makes it difficult to achieve evenly distributed immunostaining and good flatmounting. Furthermore, note that the irides or choroids are never actually grasped with forceps or directly handled during the dissection procedure. The first point of direct contact with these tissues is when they are lifted with the fine paintbrush or beaver blade and transferred into PBS. 

Protocol for Immunohistochemical Staining of Iris and Choroidal Tissue Wholemounts

Single Immunostaining of Tissue Wholemounts.

Tissue pieces are incubated in small sealed glass vials containing prewarmed 20 mM EDTA tetrasodium for 30 minutes at 37°C. The tissue in each vial is then quickly rinsed in PBS three times at 10 minutes per rinse. Increased permeabilization of the tissue is achieved by replacing PBS in the vials with a 0.1% solution of Tween-20 (or similar detergent) in PBS plus 1% bovine serum albumin (BSA; wt/vol; 20 minutes at room temperature [RT], 22°C). 

Tissue pieces are allocated to individual wells in a flat-bottomed 24-multiwell plate (Iwaki Glass, Tokyo, Japan) corresponding to the number of primary antibodies to be used in the experiment. Incubation of tissue with primary antibodies (e.g., mouse anti-rat monoclonal) is carried out at 4°C (overnight incubation) or at RT for 1 hour. All antibodies are diluted in PBS containing 1% BSA; 150 to 200 μl is placed in each well. Between each incubation, samples are thoroughly washed (three changes of PBS, 5–10 minutes each). Solutions are carefully removed from the wells and replaced using disposable plastic or glass pipettes. It is useful to place the 24-multiwell plate on a dark background to aid in visualization of the tissue pieces in the individual wells. 

A directly conjugated secondary antibody (e.g., sheep anti-mouse horseradish peroxidase [HRP ]) or a biotinylated secondary antibody is applied for 40 minutes (RT). The secondary antibodies are also diluted in PBS containing 1% BSA with the addition of 10% vol/vol normal rat serum (if species of tissue under study is rat). If biotinylated secondary antibodies are chosen, this is followed by streptavidin conjugate (e.g., streptavidin-HRP). 

Chromogen development is performed according to individual protocols (see Appendix). 

Development of horseradish peroxidase as a red reaction product with 3-amino-9-ethylcarbazole (AEC; Sigma, St. Louis, MO) takes place for approximately 10 minutes at 37°C in a concentration of 0.2 mg AEC per milliliter of 5 mM prewarmed acetate buffer (Appendix). Visualization of labeled cells as a brown reaction product can be achieved by the use of 3,3′-diaminobenzidine (DAB). Incubation is performed for 10 to 15 minutes at RT in a concentration of 12 mg DAB per milliliter PBS . A blue reaction product can be obtained by using alkaline phosphatase–conjugated secondary antibodies and visualization of labeled cells using 0.25 mg fast blue BB base (Sigma; F-0125) per milliliter of prewarmed Tris buffer and 0.125 mg naphthol AS-MX phosphate (Sigma; N 5000) per milliliter, for approximately 20 minutes at 37°C . Reactions are stopped by replacing substrate solutions with several changes of PBS and a final rinse in distilled water. 

A range of chromogens can be selected, depending on the color of reaction product desired. With albino tissues, a range such as DAB (brown), AEC (red), or alkaline phosphatase-fast blue (blue) may be chosen. However, with pigmented tissues, brown and red chromogens are not recommended. Pigment (melanin) can be removed by bleaching before histochemical or immunohistochemical staining, but appropriate controls should be performed to ensure the bleaching process does not compromise antigen preservation in the tissue. Immunofluorescence methods may also be chosen when pigmented tissues are investigated; however, care should be taken that sufficient transillumination is still possible and that tissue autofluorescence is evaluated in negative controls. 

Mounting of Tissue Pieces.

When chromogen development with AEC or fast blue BB has been performed, tissues are placed on subbed (chrome alum and gelatin) microscope slides in a drop of water-based mounting medium (ImmunoMount; Shandon, Pittsburgh, PA). Solvent-based mounting media destroys AEC and fast blue staining and can lead the examiner to suspect negative results. If DAB is chosen as a chromogen, tissues are placed in clean glass vials and dehydrated through graded alcohols and xylene before they are placed in a drop of mounting medium Distrene-80 Plasticizer and Xylene (DPX; BDH, Poole, England) on a microscope slide. Tissues are gently spread flat using fine paint brushes before the coverslips are applied. Small brass or lead weights are placed on the coverslips overnight to flatten samples. When tissues are mounted with water-based mounting medium, the edges of the coverslips are sealed with fingernail polish to prevent drying. 

Double Immunostaining of Wholemount Tissue.

The protocol for double immunohistochemical staining is based on the work of Claassen et al. ⁷ and is identical with that described, until the step subsequent to incubation in the directly conjugated secondary antibody (recognizing primary antibody 1). Tissues are washed (three times in PBS, 5 minutes each) before incubation for 3 hours at RT with a biotinylated second primary antibody (antibody 2), diluted in PBS containing 1% BSA and 10% normal mouse serum (if the primary antibody is of mouse origin). Tissues are then incubated for 40 minutes at RT in streptavidin-alkaline phosphatase diluted in 1% BSA in PBS. The alkaline phosphatase chromogen (associated with primary antibody 2) is developed first, using fast blue BB for 20 minutes at 37°C (Appendix). Samples are then washed thoroughly, and a second chromogen is used to visualize the HRP–conjugated secondary antibody bound to primary antibody 1. The author routinely uses AEC (10 minutes, 37°C) for this chromogen, because the red reaction product contrasts well with the blue chromogen. Double-labeled cells appear purple or a mixture of red and blue if the primary antibodies are localized to different cell compartments. A final wash in PBS and distilled water (5 minutes each) is followed by application of coverslips with water-based medium (ImmunoMount; Shandon), as has been described. 

As with most techniques involving multiple steps, there are a number of stages at which problems can occur that will compromise the final results. To identify the likely cause of problems, a troubleshooting guide is included in Table 1 . 

Discussion

The use of tissue wholemounts is a widely accepted tool in the fields of retinal neurobiology, in which topographic changes in density and morphology of cellular components of the retina underpin our understanding of retinal function. Tissue wholemounts have also been used to a limited extent to study the biology of Langerhans’ cells (dendritic cells) in the corneal limbus. However, the method of dissection, preparation and staining of iris, ciliary body, and choroid wholemounts has not to our knowledge been thoroughly documented. The latter methods have been valuable in ongoing investigations of dendritic cells and resident tissue macrophages in the uveal tract and in studies of the dynamics and characteristics of the cellular infiltrates in various forms of uveitis. ^{1 2 3 4 5 6} This approach consistently provides the observer with unparalleled views and appreciation of the networks and patterns of individual cellular, neural, and vascular components within these tissues that could not be obtained using conventional sections. Furthermore, focal events (such as inflammation) that could be easily missed in conventional histologic or frozen sections are readily detected in the plan or birds-eye view afforded by using whole tissue mounts. 

Recently, several investigators have begun to explore iris wholemount methods in a wide variety of studies, ^{8 9 10} and many more have expressed an interest in using the techniques but have experienced a range of technical difficulties. It is hoped the methods described in this article provide sufficient detail for investigators to obtain maximum benefit from this alternative approach and therefore enhance the range of techniques available in their investigations of ocular immunobiology. 

Appendix 1

Substrates for Chromogen Development

A. Visualization of HRP Using AEC

1.  
   Make a 5-mM acetate buffer (0.41 g Na acetate in 100 ml distilled water[ DW]). Adjust pH to 5.0 with glacial acetic acid.
2.  
   Warm 19 ml of this buffer to 37°C.
3.  
   Dissolve 4 mg AEC in 0.5 ml N,N-dimethyl formamide. Add acetate buffer while stirring.
4.  
   Add 7 μl of 30% H₂O₂ solution (stored in a refrigerator in a light-tight container that is kept tightly closed). The resultant solution can be filtered through a Millipore filter (Bedford, MA). If filtered, discard the first 1 ml.
5.  
   Fill wells of microtiter plate with approximately 0.5 ml of this solution. Incubate at 37°C for 10 minutes.
6.  
   Stop reaction by rinsing in several changes of PBS.
7.  
   Rinse once in DW and mount in water-based mounting medium (author uses Immunomount; Shandon).

B. Visualization of HRP Using DAB

1.  
   Thaw 1 ml aliquot of DAB solution (12 mg DDB per milliliter PBS) and add 9 ml PBS. Just before use, add 5.5 μl H₂O₂.
2.  
   Place DAB solution in syringe fitted with a filter (Millipore; Bedford, MA). Note: Test DAB solution on HRP containing secondary antibody or streptavidin-HRP. A dark brown stain should appear immediately if the stain is working.
3.  
   Fill wells with approximately 0.5 ml of chromogen. Incubate at RT for 10 to 15 minutes.
4.  
   Wash in PBS to stop reaction.

C. Visualization of Alkaline Phosphatase Using Naphthol AS MX Phosphate and Fast Blue BB

1.  
   Make Tris buffer (1.21 g Tris in 100 ml DW). Adjust pH to 8.5 with concentrated HCl.
2.  
   Warm 40 ml to 37°C.
3.  
   Dissolve 5 mg naphthol AS-MX phospate ⁽¹⁾ in 250 μl N,N-dimethyl formamide. Add to warm Tris buffer while stirring.
4.  
   Add 250 μl 2M HCl (0.86 ml concentrated HCl in 5 ml DW) to 10 mg fast blue BB ⁽²⁾ , swirl, and add 250 μl fresh 4% sodium nitrite ⁽³⁾ solution. (0.4 g/10 ml DW). Mix, uncapped 1 to 2 minutes and add by drops to Tris buffer on stirrer.
5.  
   Dissolve 10 mg levamisole in 2 ml Tris buffer ⁽⁴⁾ and add to warm Tris solution.
6.  
   Ensure mixture is at 37°C and filter through 0.2/0.45 μm filter (Millipore) directly onto tissues.
7.  
   Incubate in substrate solution for 20 minutes at 37°C.
8.  
   Stop reaction by replacing substrate solution with 2 to 3 changes of PBS.

Items designated with superiors (1) through (4) can be weighed out in aliquots in advance. 

 Supported by the National Health and Medical Research Council.

 Submitted for publication September 15, 1999; revised January 18, 2000; accepted February 2, 2000.

 Commercial relationships policy: N.

 

Figure 1.

 

(1) Intact enucleated albino rat eye as seen in the binocular microscope during the dissection procedure. (2) Rat eye, illustrating the position of the initial equatorial incision behind the limbus (arrows). (3) Same eye, after completion of the equatorial cut, is separated into anterior and posterior segments. Y-shaped sutures are visible on the lens. (4) Anterior segment after removal of the lens as seen when viewed from behind. (5) The anterior segment after it has been divided into three pie-shaped pieces. (6) One segment of isolated iris–ciliary body after its removal from the cornea–sclera. Note there are no blood-filled vessels visible. This acts as evidence of a successful perfusion. (7) Frontal view of the intact posterior segment. (8) An isolated portion (approximately one third) of the posterior segment that has had the neural retina (NR) removed (left) from the rest of the globe (right) which consists of the RPE (not visible in this albino eye), choroid, and sclera. (9) Thin delicate choroid (C) (left) after its separation from the inner surface of the sclera (S) (right).

Table 1.

 

Troubleshooting Guide

Problem	Probable Cause	Remedy
Perfusion
Cannula pierces aorta	Cannula is too sharp	Blunt or round off edges with file or sandpaper.
Poor perfusion speed	Cannula tip is compressed against wall of aortic arch, reducing flow	Place tip of cannula in ascending aorta and avoid kinking the aorta.
Poor perfusion (no muscular spasms in hindlimbs, forelimbs, or tail)	Excessive time delay between opening chest cavity and cannulation. Death and hemostasis results in poor perfusion	Avoid delays and have all necessary instruments at hand.
	Blockage in tubing	Remove all air bubbles from tubing.
Lungs rapidly turn white	Cannula mistakenly placed in right ventricle, thus perfusing pulmonary vasculature	Remove cannula and reposition in left ventricle.
Dissection
Tissue wholemounts will not flatten	Tissues allowed to dry out in folded position	Always keep tissue moist during dissection.
Iris adherent to cornea and retina adherent to choroid	Overfixation	Dissect eyes immediately after perfusions. Do not store in fixative for long periods.
Processing
Loss of tissue pieces	Lost during pipetting	Take care when pipetting to avoid sucking up tissue pieces.
	Pieces adherent to side of the well	Pipetting fluid down the side of wells dislodges adherent pieces. Reduce number of air bubbles in wells. Count pieces before beginning and check wells regularly. Placing multiwell plates on a sheet of black paper aids in visualizing the tissues. Pipette waste fluids into a beaker placed on black background, to aid in detecting inadvertently expelled tissue pieces.
Wells dry out	Insufficient fluid in each well	150–200 μl of fluid should be adequate for each well.
	Evaporation of fluids	The lids of plates placed in refrigerator overnight must be wrapped tightly with plastic sealant or wrapping.
Tissues damaged or folded	Excessive handling and touching during dissection or by pipette during changing of reagents	Use plastic disposable pipette tip on the end of glass transfer pipette. Pipette fluids from side of the wells, avoiding tissue.
	Drying out of wells between fluid changes	Minimize delays between fluid replacements.
Staining Results
Excessive background staining	Inadequate fixation	Fix for longer period.
	Poor perfusion (i.e., pseudoperoxidase staining of RBCs in vessels)	Repeat experiment and ensure good perfusion.
	Concentration of primary or secondary antibody too high	Titrate primary and secondary antibodies.
	Absence of blocking antibody in secondary step	Include 10% serum from species under investigated in secondary antibody solution.
	Overdevelopment of chromogen	Watch development of chromogen and halt reaction sooner.
Unexpected staining pattern	Inadvertent transfer of tissue between wells occurring after incubation with primary antibody	Change pipette tips between wells. Care should be taken to avoid contaminating adjoining wells that have different primary antibodies.
Patchy staining (some areas devoid of stained cells)	Tissue has been folded during incubations or is in contact with air bubbles	Examine the wells with a dissecting microscope or magnifying lens when changing solutions to ensure tissues are floating in solutions.
Difficult to focus on cells in microscope	Wholemounts not flattened sufficiently	Increase weight on coverslips.
	Excess mounting media	Remove ciliary body and mount separately.
Fluorescent labeled cells appear dull	Incubations not performed in dark	Cover multiwell plate with tinfoil or keep in a black box.
No staining	Inappropriate primary or secondary antibody	Check protocol and ensure primary monoclonal antibody is raised to recognize the species under investigation.
	No activity in primary antibody	Check that the secondary antibody recognizes the species the primary antibody was raised in.
	No H2O2 added to chromogen, or H2O2 is old and out of date	Check activity of primary antibody on positive control frozen sections before staining wholemounts.
	Underdevelopment of chromogen	Prewarm chromogen to 37°C.
Stain deposits throughout tissue	Failure to filter chromogen or centrifuge fluoresceinated antibodies	Pass through filter before use, or centrifuge.

The author thanks the numerous students and research assistants who have assisted over several years in refining the methods described in this article.