Methods for Imaging Inflammation and Transendothelial Migration in vivo and ex vivo (2024)

Curr Protoc. Author manuscript; available in PMC 2024 Apr 1.

Published in final edited form as:

PMCID: PMC10309184

NIHMSID: NIHMS1886781

Vivienne Fang,^*# Maureen E. Haynes,^*# Vanessa Hayashi,^* Erika Arias,^* Jeremy A. Lavine,^** David P. Sullivan,^* and William A. Muller^*

Author information Copyright and License information PMC Disclaimer

Starting materials:

Mice (we have tested FVB, C57BL/6, 129S backgrounds)

Carrier solution (see recipes in Reagents and Solutions)

Croton oil mixture (see recipes in Reagents and Solutions)

Nair^™ hair-removal lotion

DPBS, 1X without calcium & magnesium (CORNING, lot 26321007)

Fixing solution (see recipe in Reagents and Solutions)

Bovine serum albumin (BSA) solution (see recipe in Reagents and Solutions)

Permeabilization buffer (see recipe in Reagents and Solutions)

Normal goat serum (NGS) (Jackson, cat. no. 005-000-121)

Normal mouse serum (NMS) (Jackson, cat. no. 015-000-120)

Anti-PECAM 2H8 (Millipore Sigma, cat. no. MAB1398Z)

Anti-S100A9 (Abcam, cat. no. ab105472)

Fluorophore-conjugated goat anti-Armenian hamster secondary

Fluorophore-conjugated goat anti-rat secondary

FluorSave mounting reagent (Calbiochem, cat. no. 345789–20L)

Hardware and instruments:

1.5 mL microfuge tubes

Isoflurane anesthetic apparatus

Heating pad

CO₂ tank with mouse euthanasia apparatus

Cotton swabs

Dissection scissors and tweezers

15 mL conical tubes

12 well flat bottom plate

Plate shaker

100 mm culture dish

Dissection microscope

24 well flat bottom plate

Aluminum foil

Glass microscope slides

Coverslips #1.5, 22mmx22mm

Spinning disk confocal microscope

Day 1

1. Prepare the carrier and croton oil mixtures in two 1.5mL microfuge tubes.

2. Set the air flow rate to 2 L/min and anesthetize the mice with isoflurane. Wait for the mouse’s breathing to slow down enough to indicate a sufficient plane of anesthesia.

   This procedure does not have to be performed under anesthesia if the investigator is comfortable “scruffing” the mouse or if there is a gentle way of immobilizing the mouse’s head. For consistency, the croton oil and carrier mixtures must be applied evenly and completely, which is easier when the mouse’s head is immobilized.

3. Pipette 10 μL of carrier mixture onto the back of the left ear and 10 μL of the same carrier mixture onto the front side.

4. Pipette 10μL of croton oil onto the front and back of the right ear. Repeat for all mice.

5. Leave mice in cage with access to a heating pad for 5 hours.

6. Sacrifice mice with CO₂ asphyxiation and cervical dislocation.

7. Rub Nair on both sides of the ears using a cotton swab, leave for a minute and wipe off with disposable wipes to remove hair. Cut the ears off the mouse. Pour 12 mL of 1X DPBS into each of two 15 mL conical tubes, and place an ear into each tube, keeping track of which ear received which treatment. Shake the tubes to wash the Nair off the ears.

   The bubbles will automatically bring the ears to the surface of DPBS, so they can be easily removed with tweezers. The DPBS can be reused for more mice, but it is recommended to pour fresh DPBS when it is murky.

8. Place the ears into a labeled 12 well flat bottom plate. Fill each of the wells with 1mL fresh fixing solution. Leave the plates on a shaker for 0.5–1hr at room temperature.

9. Add 1X DPBS to a 100 mm culture dish. Temporarily transfer 1 ear at a time to this 100 mm dish and place under dissection microscope to zoom in on the cut edge of the ear. Separate the front and back sides of the ear by gripping each side at the cut edge with fine forceps, and then slowly pulling them apart. Keep track of which side is the epithelial side and which is the side with interstitial tissue / endothelium.

10. Making sure the interstitial tissue/endothelium side of the ear is facing down, place each ear half individually in a 12 well plate with fresh fixing solution. Leave the plate on a shaker overnight at 4°C.

    The epithelial (outer side is hydrophobic/waxy) so the ear halves may float with the interstitial above the fixative if not placed interstitial side down.

Day 2

11. Wash ears with 1X DPBS 10 min × 3 times at room temperature.

12. Place each half-ear in the same orientation as before into a 24 well plate with 500 mL permeabilization buffer in each well. Leave to permeabilize on a shaker overnight at 4°C.

Day 3

13. Wash ears with 1X DPBS 10 min × 3 times at room temperature.

14. Dilute anti-PECAM 2H8 and anti-S100A9 in a solution of 2.5% bovine serum albumin (BSA) + 5% normal goat serum (NGS). Add 250 mL antibody mixture into each well of a 24 well plate. Place each half-ear in the same orientation as before in the wells. Leave the plate on a shaker overnight at 4°C.

    Recommended concentrations are 9 μg/ml anti-PECAM and 2 μg/ml anti-S100A9, which corresponds to dilutions of 1:200 for anti-PECAM 2H8 and 1:500 for anti-S100A9 from the commercial sources stated above.

Day 4

15. Wash ears with 1X DPBS 10 min × 3 times at room temperature.

16. Dilute goat anti-Armenian hamster secondary and goat anti-rat secondary in a solution of 2.5% BSA + 5% NGS + 5% normal mouse serum. Leave this mixture for 15 min on ice to pre-absorb the secondary antibodies. Add 250 mL of the mixture into each well of a 24 well plate. Place each half-ear in the same orientation as before in the wells. Cover the plate with aluminum foil to protect the fluorophores. Leave the plate on a shaker for 4 hrs at room temperature.

17. Wash ears with 1X DPBS 10 min × 3 times at room temperature.

18. Mount specimens on glass microscope slides. Place two drops of FluorSave mounting reagent onto the slide, and place the half-ear onto the slide with the epithelial side facing down. Add as many drops of FluorSave as needed and place coverslip on top. Allow them to dry in a dark place for at least 1 hr at room temperature.

    Place the coverslip delicately so that as few bubbles as possible get trapped. Depending on the shape of the half-ears, they might need to be trimmed so that they can lay flat on the slide. It is recommended to use a generous amount of FluorSave reagent, because it has a tendency to shrink when dry.

19. Store slides in a slide holder to maintain them protected from light and store at 4°C until imaging.

    Slides can be stored for several months without significantly decreasing the quality of the images, but it is still recommended to image them soon after mounting.

Imaging

20. Turn on the spinning disk confocal microscope. Place the slide on the mechanical stage and add a drop of immersion oil on top of the coverslips. Adjust the position of the slide so that the objective lens (40X oil) is directly above the ear that received carrier mixture. Lower the objective until it barely touches the drop of oil.

    Since this protocol uses fixed samples for imaging, a laser scanning confocal can be a replacement for the spinning disk confocal.

21. Focus the image until the blood vessels are visible in the goat anti-Armenian hamster secondary channel.

22. Switch to the goat anti-rat secondary channel, and scan to verify that there are few to no leukocytes (0–2 per field).

    The ears that received carrier mixture should not have inflammation, otherwise they cannot serve as control conditions for the ears on which acute inflammation was triggered using croton oil mixture.

23. Adjust the position of the slide so that the objective lens is directly above the ear that received croton oil mixture. Lower the objective and focus the image as in step 2.

24. Scan the sample for blood vessels, whose cell-to-cell junctions have a cobblestone pattern, and that have a diameter between 15μm and 50μm, ideally between 30μm and 50μm.

    These are most likely to be post-capillary venules, where neutrophils extravasate. Avoid vessels that have highly elongated, thin endothelial cells because these are likely arteries.

25. Switch the laser to the goat anti-rat secondary channel. Take a Z-stack if there are 10–30 leukocytes in the field.

26. Repeat steps 5 and 6 for 10 or more fields in each ear that received croton oil mixture.

27. Repeat entire imaging process for all the experimental conditions.

Counting leukocytes

28. On each picture, look at the Z-plane and score how many leukocytes are inside and outside the vessels (area within 50mm from a venule). If there are leukocytes that are not fully inside nor outside, count them separately.

    Avoid counting leukocytes inside the vessels that are not in contact with the walls of the vessels, as much as possible. Avoid counting leukocytes that are outside of vessels with low amount of fluorescence, because these are likely to be resident macrophages.

29. Calculate the percent TEM per image. Divide the number of leukocytes that were outside the vessels by the total amount of leukocytes in the given image. Repeat this for all the images.

30. Calculate the average percent TEM. Take the average of the values from step 2. Compare the average percent TEM between/among experimental conditions and perform statistical tests as needed.

    If the average percent TEM in an experimental condition is lower than the control condition, it means that TEM was blocked to a given extent by the condition the mouse was exposed to.

Table 1.

Troubleshooting Guide for Croton Oil Dermatitis

Starting materials:

Mice with genetically-encoded fluorescent leukocytes, wild-type or knockouts, with or without a treatment of interest (see Fluorescent Mice section below)

DPBS, 1X without calcium & magnesium (CORNING, lot 26321007)

Recombinant mouse interleukin-1β (IL-1β) (R&D Systems, 401-ML-005/CF)

Ketamine hydrochloride (Covetrus, NDC# 11695-0703-1)

Xylazine (Akorn Inc., NDC# 59399-110-20)

Anti-PECAM antibody conjugated to a fluorophore (EMD Millipore, CBL1337 clone 390, Rat anti-mouse, non-blocking)

Nair^™ hair-removal lotion

Perfusion buffer (see recipe in Reagents and Solutions)

Tyrode’s Salts (Millipore Sigma, T2145–10X1L)

Sodium Bicarbonate (Millipore Sigma, T2145)

Hardware and Instruments:

Plexiglas platform with glass window

Silicone stage with quartz pedestal

Dissecting microscope (Leica or equivalent)

Fine forceps and scissors

Threaded needle

Suture needles bent into an “L” shape

Dissecting pins

Buffer warmer

Cotton tip applicators

Heating pad

Syringe pump

Confocal microscope (Nikon or equivalent) with 20× water-immersion objective

Starting materials:

SYLGARD 184 Silicone Elastomer Kit (Dow Corning # 4019862)

Clear fused quartz ground and polished disc, 1×0.25in (Technical Glass Products #1X0.25)

6-well tissue culture dish

Modeling clay

Hardware and instruments:

Vacuum chamber

Dissecting light

Ideal Fields

One of the best ways to ensure consistent results and striking movies is to pick consistent fields to observe and record. The ideal venule will:

1. be largely straight and not tortuous for at least 150 μm in length,

2. be 30–50 μm in diameter,

3. be near the surface/closer to the objective,

4. have consistent flow without any pulsating or occasional pausing, and

5. have signs of inflammation including adherent and rolling leukocytes and some leukocytes already out in tissue.

Although it is common to have to compromise on some these details, it is important to understand how they may affect the results. It is difficult to measure velocities in tortuous vessels and they often have abnormally high adhesion. Narrower vessels and vessels with inconsistent flow usually have slower flow rates and often dramatically increased adhesion and slower rolling. In addition, vessels with intermittent flow will often cease flowing by the end of the recording, making the entire effort fruitless. Signs of active inflammation is often a good indicator of ongoing recruitment and TEM. However, one must be careful not to bias the results by only choosing ‘hot’ areas. In almost every mouse, there are regions of hyper- and hypo-inflammation, possibly due to damage during tissue preparation or exposure to the stimulus. Where rolling and adhesion are to be examined, it is more important to compare vessels of similar size and tortuosity. Where TEM is to be examined, extra attention should be given to finding venules that have a modest amount of active inflammation, to avoid hyperactive areas (obviously little TEM can occur where there is no rolling or adhesion.)

Drifting field and tenting

It is somewhat common for the field to drift (move in x and y) slightly over the course of the recording due to changes in the muscle tension and relaxation. Small drifts can be corrected in the image processing software to align the images together. If given the option, align the time points according to the vessel fluorescence channel, because the structures are more consistent than the leukocyte channel. If the field moves too much, longer recordings may become impossible. Typically, fields are more stable at the periphery of the tissue near where it has been pinned. Unfortunately, these regions are less likely to have appropriate venules to record. Drifting can be limited somewhat by using more pins to secure the tissue, especially proximal to the body. Take care when adding pins in this region though, it is easy to apply too much tension to the major vessels that feed and drain tissue thus affecting the flow rates to the entire tissue. Another common issue is tenting or drifting in the z direction. This occurs when fluid slowly accumulates under the tissue pushing it up and moving it out of focus. Slow changes can be corrected for by adjusting the z-focus manually (if the scope allows). Tenting can typically be avoided by making sure that the tissue exits the body parallel to the silicone mount and pedestal, and that it is well secured. Keeping the flow rate of the perfusion buffer slow but steady will help keep it from accumulating beneath the tissue.

Tissue drying out

Typical recordings of TEM can go for 15 minutes or longer. During this time, the perfusion buffer running down the objective will hydrate the field of view. However, depending on subtleties of the tissue and silicone support, the distal regions of the tissue may not receive adequate flow to remain viable. After finding a view of interest, take note of the hydration state distal regions of the tissue to make sure they are receiving enough buffer to remain hydrated. Ideally, much of the tissue will be clearly covered with buffer. To prevent the tissue from drying out and ruining potentially ideal fields, the flow rate of the perfusion buffer can be increased. This issue is more common for regions visualized near the periphery and less so in nearer the center of the tissue. If this is not possible alter the flow rate or change positions, the tissue may be hydrated manually by gently pipetting a small amount of warm perfusion buffer directly onto the tissue. Take care to apply it between images and avoid touching the lens or tissue.

Fluorescent mice

To avoid the use of labeling antibodies that might interfere with leukocyte or endothelial cell function, it is often easiest to use mice that express fluorescent proteins in the specific subset of leukocyte one wishes to observe. For example, GFP inserted into the LysM locus adequately labels all myelomonocytic cells (neutrophils, monocytes, and macrophages), although neutrophils are typically brighter than monocytes while macrophages are somewhat dim, more ramified, and within the tissue rather than intravascular. Along these lines, it is possible (and convenient) to use mice whose leukocytes have been rendered fluorescent. Mice with genetically encoded fluorescent leukocytes are available with virtually any leukocyte subtype labeled. These can be bred to your mouse of interest or, assuming histocompatability, provided via a standard adoptive bone marrow transfer.

Table 2.

Troubleshooting Guide for Intravital Microscopy of Mouse Cremaster Muscle

Starting materials:

Genetically modified mice with fluorescent leukocytes, such as LysM^GFP, Catchup^IVM or Ccr2^RFP mice

Anti-PECAM antibody conjugated to a fluorophore (EMD Millipore,CBL1337 clone 390, Rat anti-mouse, non-blocking)

DPBS, 1X without calcium & magnesium (CORNING, lot 26321007)

4% paraformaldehyde solution (see recipe in Reagents and Solutions)

Hardware and Instruments:

30 mL syringes

Fine forceps and scissors

Dissecting pins

Razors

Murine Brain Matrix (RWD; device for holding brain in place for precise slicing)

Coverslip dishes (Mattek with #1.5 coverslips)

Wide-field microscope (Nikon or equivalent) with 4X objective

Table 3.

Troubleshooting Guide for Wide-field Microscopy of the Murine Brain

Starting materials:

Avertin (see recipe in Reagents and Solutions)

70% ethanol

4% paraformaldehyde fixing solution (see recipe in Reagents and Solutions)

Sucrose

DPBS, 1x, without calcium & magnesium (Corning, #26321007)

OCT Compound (Sakura #4583)

Dry ice

Hardware and Instruments:

Dissecting board

Dissecting tools

Dissecting board pins

Sutures

1mL syringe

Luer stubs (Instech Labs #LS20)

Dissecting scope

MatTek dishes (MatTek #P35G-1.5–14-C)

Tissue-Tek Cryomold 15×15×5mm (Fisher Scientific, #NC9511236)

22×22mm cover slips (Fisher Scientific, #50-143-780)

Razor blades

Cryostat blades

Brush

Slides

Cryostat for tissue sectioning

Widefield microscope

FIJI software

Optional materials:

Labeling antibodies (e.g. 390 to label endothelial cell junctions)

Brain matrix (RWD)

Please see Figure 8 for a general workflow for imaging the lungs ex vivo, the details of which are described below.

Removing the lungs:

1. Induce inflammation or perform the procedure necessitated by the experiment along with any appropriate controls.

2. Prior to euthanization, inject any needed live antibody labels. Make sure that these are fixable.

   Typically, 30 minutes prior to euthanasia, we label PECAM-1 with non-blocking 390 antibody and 2,000,000MW dextran.

3. Inject the mouse with an overdose injection of Avertin (800ul-1ml of Avertin, prepared as noted previously) to achieve deep anesthesia.

   This can be performed by any IACUC accepted procedure, however because many anesthesia methods are inhaled, you must be mindful of potential for your method to cause further damage/inflammation in the lungs.

4. Once the mouse is in a deep plane of anesthesia (unresponsive to toe pinch), you can proceed with the lung removal.

5. To remove the lungs, place the mouse flat on a dissecting board on its back, pinning all four limbs to the board using pins.

6. Use 70% ethanol to wet the hair on the top of the mouse, focusing on the abdomen and the head. This is not to sanitize or clean the mouse but rather for fur control (to avoid getting fur in the sample).

7. Gross dissect the skin away from the abdomen by gently pulling up on the skin on the abdomen with blunt forceps and making a small incision with blunt-tipped scissors. Insert the scissors in the cut, below the skin but above the chest plate. Open and close the scissors, gently moving them up the abdomen until you’ve bluntly dissected the skin away from the chest plate up to the jaw.

8. Using the blunt tipped scissors, cut away the loose skin covering the chest and bottom of the jaw.

9. Underneath the jaw of the mouse, there will be two pieces of white-pink tissue (salivary glands). Pull these two apart or remove them completely, exposing the trachea.

10. Remove the chest wall by cutting away the diaphragm and up the left and right side of the ribs. This will expose the lungs.

11. Carefully dissect away the protective tissue around the trachea, exposing the horizontal rings of cartilage. You should dissect away the tissue both ventrally and dorsally.

12. Thread a 10cm piece of suture under the trachea, leaving equal visible trachea space above and below the suture.

13. Using small, sharp scissors, make a small (~1mm) cut in between the rings of cartilage of the trachea.

    To make this easier, you can gently tug up on the suture, pulling up and exposing more of the trachea from the body. However, you want the trachea to remain in one piece connected to the lungs.

14. Insert the Luer stub into the incision and tightly tie the suture around the Luer stub and trachea with a single square knot that can be undone.

15. Slowly instill the lung with up to 700ul of fixative using a 1ml slip tip syringe.

    The volume of murine lungs is approximately 1ml, depending on the size of the mouse. You want to inflate the lungs to retain the integrity of the alveolar structure, but not overinflate them. We fix with 4% PFA.

16. Remove the syringe and Luer stub while simultaneously tightening the suture to keep the fixative inside the lungs. Tie off the suture with a box knot.

17. Holding the trachea up with the suture (gently pulling away from the body), cut at the top of the trachea, then dissect the lungs and heart away from the body.

18. Remove the lungs/heart together with the suture. Place in a container filled with fixative.

19. At this point, you can either quick-fix or fully fix. If you are quick-fixing, leave the lungs in the fixative for 30 minutes then move to imaging. If you are fully fixing, leave in fixative overnight. The next day, transfer the lung to 15% sucrose in PBS. Once the lungs have sunk to the bottom in the 15% sucrose (usually 1–2 days), transfer them to 30% sucrose until they sink (1–2 days).

20. Once the lungs are ready for imaging, you can image whole lungs, thick sections, or thin sections. You can also split the lungs up into left and right lungs or individual lobes for different imaging methods.

    For confocal imaging, we typically use whole lung lobes. For widefield or epifluorescence, we typically use thick or thin sections.

For whole lung imaging:

21. Remove the lungs from fixative. Dissect away any blood clots or excess tissue from the outside of the lungs.

22. Using sharp scissors, cut off the whole lobe that you plan to image. Be careful to not cause any trauma on the outside of the lungs. This can be ensured by holding the heart with forceps rather than the lung itself.

23. Under the dissecting microscope, pop any surface bubbles and clear the top of the lobe of blood and tissue as much as possible. Bubbles will continue to rise to the surface as a result of the physiology of the lung, just do your best to pop surface bubbles.

24. Place the lobe, viewing side down, in a MatTek dish. Hydrate with PBS and place a cover slip on top.

25. The lung is now prepared for imaging.

For thick sectioning:

26. Remove the lungs from fixative. Dissect away any blood clots or excess tissue from the outside of the lungs.

27. Using sharp scissors, cut off the whole lobe that you plan to image.

28. Place the lobe in a Tissue-Tek Cryomold and either quick or slow freeze the lobe. To quick-freeze the lobe, place the cryomold on dry ice (5–10 minute). To slowly freeze the lobe, place the cryomold in a −20C freezer until frozen (30–60mins).

29. Remove the lobe and section in 1mm slices with a razor blade. This can be done by eye or in a brain matrix for sectioning. Using a brain matrix works best on the left lobe of the mouse.

30. Place the section in a MatTek dish in 3–4 drops of PBS. Under dissecting microscope, pop any surface bubbles and place a cover slip on top.

31. The lung is now prepared for imaging.

For thin sectioning:

32. Remove the lungs from fixative. Dissect away any blood clots or excess tissue from the outside of the lungs.

33. Using sharp scissors, cut off the whole lobe that you plan to image.

34. Place a dot of OCT in a 15×15×5mm Tissue-Tek Cryomold. Wait until the OCT has settled at the bottom, and all bubbles have popped. The OCT should fully coat the bottom of the Cryomold.

35. Place the lung lobe you plan to image on the OCT in the cryomold. Cover the lung with more OCT until it is fully submerged. Allow the OCT to settle and all the bubbles to pop (2–3 mins).

36. Place the cryomold on dry ice.

37. Once frozen, remove the frozen block from the mold and place inside a cryostat set to 18C.

38. With a razor blade, cut off the edges of the OCT that don’t contain lung tissue.

39. Using the Cryostat, section the lung tissue into 10–15um sections and place on a slide. Keep the slides uncovered on dry ice.

40. Rinse the OCT off by placing 2–3 drops of PBS on the slide for 5 minutes before placing a cover slip on the slide.

41. The slides are now ready to image.

Table 4.

Troubleshooting Guide for Imaging inflammation in the murine lungs (ex-vivo)

Starting materials:

Ketamine/Xylazine concoction (see recipes in Reagents and Solutions)

Ketamine hydrochloride (Covetrus, NDC# 11695-0703-1)

Xylazine (Akorn Inc., NDC# 59399-110-20)

Proparacaine hydrochloride eye drops

Phenylephrine hydrochloride eye drops

Tropicamide eyedrops

DPBS, 1X without calcium & magnesium (CORNING, lot 26321007)

4% paraformaldehyde fixing solution (see recipe in Reagents and Solutions)

Blocking solution (see recipe in Reagents and Solutions)

Washing solution (see recipe in Reagents and Solutions)

ProLong^™ Gold Antifade Mountant (Thermo Fisher; catalog # P36930)

Recombinant Mouse CCL2/JE/MCP-1 Protein (R&D Systems Incl; catalog # 479-JE-050/CF)

Rabbit Anti-S100A9 antibody [2B10] (Abcam; ab105472) to detect neutrophils

Rabbit Anti-IBA1 antibody (Wako, 019–19741) to detect macrophages

Goat Anti-Collagen IV antibody (Abcam; ab19808) to detect vessels

Goat Anti-CD31 antibody (R&D Systems, AF3628) to detect vessels

AffiniPure Goat Anti-Rat IgG (Jackson; Lot # 112-005-167) – conjugated in house

Alexa Fluor^® 647 AffiniPure Goat Anti-Rabbit IgG (Jackson; Lot # 111-605-144)

Hardware and Instruments:

Ear punch tool

Scale

1 mL syringe

30G needle

5 mL syringe

Scissors

Model 901 Removable Needle Syringe, 10 μL capacity (Hamilton, Ref # 7648–01)

Small Hub Removable Needle, 32 gauge, 0.4 inch, point style 3 (Hamilton, Ref # 7803–04)

CO2 tank with mouse euthanasia apparatus

Carcass disposal bag

Curved forceps (Fine Science Tools; No. 91117–10)

Light microscope

Ring forceps (Fine Science Tools; No. 11103–09)

Spring scissors (Fine Science Tools; No. 15003–08)

Straight forceps (x2; Fine Science Tools; No. 11254–20)

Weigh boats

12-well plates (Thermo Scientific; Catalog # 130185)

24-well plates (Thermo Scientific; Catalog # 130186)

Orbital shaker

3 mL transfer pipettes (VWR; catalog # 52947–948)

3” × 1” × 1.2 mm microscope slides (VWR; catalog # 16004–370)

20 × 60 mm, #1.5 microscope cover glasses (VWR; catalog # 16004–312)

Stirring hotplate (Thermo Fisher; catalog # SP88854100)

Tagging and Sedating Mice

1. Obtain mice of interest, such as C57BL/6 or Ccr2^RFPCx3cr1^GFP mice.

2. If using multiple mice, tag to differentiate one from another. Suggestions for tagging include ear punch or ear tag.

3. Weigh mice.

4. Calculate dose of ketamine/xylazine to administer to mice based on weight. See for more information on ketamine/xylazine concoction.

5. Scruff mice and inject correct ketamine/xylazine dose intraperitoneally. Wait a few minutes until mouse is sedated.

   Mouse is sedated enough for intravitreal injections once it is immobile and its whiskers are minimally twitching.

   If mouse is not sedated enough with initial dose, boost with 25–50% of initial ketamine/xylazine injection.

   CAUTION: Injecting too much ketamine/xylazine can kill the mouse. Be careful to inject the correct dose based on weight.

6. Once mouse is sedated, inject 0.4 mL of meloxicam (1:100 meloxicam:1XDPBS mixture) subcutaneously with 30G needle attached to a 5 mL syringe. To perform a subcutaneous injection, place mouse on a flat surface, pinch and lift the skin on its back and inject into the lifted skin.

   Be sure the needle is just under the skin and not to push the needle through to the other side to prevent injury to self.

Intravitreal Injection of Inflammatory stimulus CCL2

7. Trim whiskers to prevent microscope examination disruption.

8. Scruff mouse and position its head with one eye facing up. Administer one drop of proparacaine hydrochloride to eye. Balance the drop for 2–3 seconds, then shake off. Repeat in other eye.

   Proparacaine hydrochloride is a topical anesthetic that will numb the eyes.

9. Administer phenylephrine hydrochloride eye drops (same handling technique as in step 8).

   Phenylephrine hydrochloride will dilate the pupils and retract the top eyelid back.

10. Administer tropicamide eye drops (same handling technique as in step 8).

    Tropicamide is an anti-cholinergic drug that will dilate the pupils.

11. Place mouse under light microscope on a paper towel. Position the head with the eye of interest facing up.

    Note: Lubricate eyes with artificial tears every five minutes to prevent the corneal surface from drying out. Mice lose their blink reflex when sedated.

12. With a 30G needle attached to an empty 1 mL syringe, make an incision just posterior to the sclerocorneal junction on the lateral side of the eye (Figure 11). The incision should be bevel deep and no more.

    Open in a separate window

    Figure 11.

    Schematic diagram of a mouse eye with the pupil, iris, cornea, and sclera labeled. The sclerocorneal junction, or limbus, is outlined in red, and the location for the initial incision prior to CCL2 injection is indicated with a blue star. The medial and lateral sides of the eye are also labeled; the incision should be done on the lateral side of the mouse eye. Created with BioRender.com.

    If the mouse’s eyelashes are obscuring your view of the globe, gently brush away the lashes with body of needle.

13. With a Hamilton syringe loaded with your inflammatory stimulus (CCL2, 5 ng / uL), insert the tip of the needle two (2) millimeters deep into the incision you created in step 12. Have a second person help push the plunger to inject 1 uL of CCL2.

14. Flip the mouse to its other side and repeat steps 11–13 to inject the other eye. Omit this step if you are only injecting one eye.

15. Place mouse in an empty cage on a cage warmer until the mouse wakes up.

16. Return mouse to its original cage and provide food and water as usual. Sacrifice mice to harvest eyes 24 hours after intravitreal injection.

    The amount of time to sacrifice is dependent on your experiment. 24 hours post-CCL2 injection is sufficient to visualize neutrophils and monocytes.

Enucleation and Dissection of Retina

17. At the desired time after intravitreal injection, euthanize mice per IACUC protocol.

18. Using curved forceps, enucleate the eyes: maneuver the forcep tips around and under the eye globe, pinch the optic nerve and gently pull the eye free from the socket (Figure 12). Place enucleated eyes in a dish with 1X DPBS and place under a dissecting light microscope.

    Open in a separate window

    Figure 12.

    Schematic diagram of a profile view of an eye. The eye globe and optic nerve are labeled. When enucleating the eye, it is important for the experimenter to pinch the optic nerve behind the globe and not the globe itself, as highlighted in the diagram (arrow). Created with BioRender.com.

19. Dissect globe to remove conjunctiva, extraocular muscles, and orbital fat. Stabilize globe with ring forceps and make small incision (1 mm) with spring scissors at the limbus of the eye.

20. Substitute ring forceps with straight forceps. Insert straight forcep tip into the incision from step 19 and grip the anterior portion of the eye.

21. Cut along the limbus around the globe using spring scissors in one hand, while gripping the anterior portion of the globe using straight forceps with the other hand.

22. Remove the anterior portion of the eye.

23. Separate the lens from the posterior globe.

    If the lens is adhered to the posterior cup, use forceps to grab the choroid-sclera complex, optic nerve, or any tissue that is not the retina while coaxing the lens out with straight forceps in the other hand.

24. Pinch the choroid-sclera complex with straight forceps in both hands and tear. Continue tearing at different parts of the globe until the cup is easily separable from the retina.

    Be careful not to pinch or touch the retina at any point as it is very fragile tissue.

25. Pinch the retina at the optic nerve to separate it from the cup.

Immunofluorescent Staining of Wholemount Retina

Mounting Retina onto Microscope Glass Slide

“Threshold” duplicate

1. In your “threshold” duplicate, choose a threshold strategy for your image that highlights your cells of interest. To do this, go to Image > Adjust > Threshold. Make sure “dark background” is ticked and “B&W” is selected under the second drop-down menu. In the first drop-down menu, scroll through the different threshold options until you get one that automatically highlights your cells of interest. Click “Apply” and close the window.

   It is ok if debris in the retina is also highlighted as we will remove debris in the next step. It is most important to ensure that the thresholding option you choose highlights the leukocytes/cells of interest.

2. Next we will segment out debris. Go to Analyze > Analyze Particles > 300-infinity. Make sure “add manager” is ticked in the ROI pop-up window.

   The goal of this step is to segment out areas of your image you do not want FIJI to register as cells. You can change the 300 to any particle size. This segmentation tells FIJI that you want to highlight anything greater than 300 um² (or the size you choose).

“Original” duplicate

1. Now we will overlay the debris selections from step 6 over your “original” duplicate. Go to your “original” duplicate, click “show all” in the ROI manager until you see your debris selection on your original image. Click “More” in the ROI manager > Fill.

   This will “fill” the debris of your retina with a color and remove those data so that FIJI will not register it when you count cells later. Select the fill color in the color selector in the FIJI toolbar. You can choose whichever color you wish. In the ROI manager, More > Fill to fill in the color you chose. The resulting image is the image you want to analyze.

2. Next you will measure the total area of the debris selections, as you will need to subtract this area from your selection of the whole retina later to calculate a “total retina area” to count cells in. In your ROI manager, there is a list of selections that correspond to the segmented debris. Select all points in the ROI manager, then Analyze > Measure in the menu at the top of your screen. The output is the area (um²) of the debris. Write this number down.

   You can save selections in the ROI manager by clicking More > Save.

   To ensure that “area” is the output that is measured, go to Analyze > Set measurements and make sure “area” is selected.

3. Next, we will measure the area of retina to be analyzed. This will require you to calculate the area of the entire retina (including the debris). Go to your “original” duplicate that now has the debris filled out, select the “polygon” selection tool in the FIJI toolbar, and manually outline the entire retina. By hand drawing, you can get a count of “cells per area” and compare this in a standardized way. Once you have made your selection, go to your ROI manager and click “Add” to add your selection to the manager. In the manager, click on the selection you just added, and Analyze > Measure to calculate the area of your selection in um². Note that this is not the final area. To find the total area of retina to count cells in, subtract “debris area” from the area you just measured. Write this number down.

   After making your selection, make sure you do not click anywhere else in the image because it will deselect your selection. If this happens, go to Edit > Selection > Restore Selection. Note that this will only restore your most recent selection.

   To save your selection, go to Analyze > Tools > ROI manager > More > Save

   To open or load your selection, go to Analyze > Tools > ROI manager > More > Open

4. Next find and calculate the number of leukocytes in the retina. To do this, you must tell FIJI what you consider to be a leukocyte. While making sure the retina area you selected is still highlighted, zoom in to your tissue and click the “hand/scrolling tool” button in the FIJI toolbar to move to an area of the retina with leukocytes. Go to Process > Find maxima in the menu at the top of the screen. The number of maxima is your leukocyte count.

   Make sure “preview point selection” is ticked. The bigger the prominence, the fewer points you will have. Make sure you zoom into the tissue far enough to see differences in points selected based on how you change your prominence (ie when you increase or decrease prominence). Your tissue area selected may deselect during this process, so make sure you have saved your selection area per step 9 so you can reload it.

5. Finally, quantify the number of leukocytes per retina by dividing your maxima calculated in step 10 by the final retina area calculated in step 9. This will give you # leukocytes/um² retina.

Basic Protocol 1:

Basic Protocol 2:

Basic Protocol 5:

Table 5.

Troubleshooting Guide for Leukocyte Extravasation Analysis in Retina