A clearing-free protocol for imaging intact whole adipose tissue innervation in mice

Summary Here we provide a clearing-free protocol for processing intact, whole mount subcutaneous white adipose tissue (scWAT) for immunofluorescence as an alternative to current clearing-based approaches. We use a combination of Z-depth reduction and autofluorescence quenching techniques to fluorescently label, image, and quantify adipose tissue innervation effectively throughout intact mouse tissues without the need for optical clearing or light sheet microscopy. This protocol has been optimized and validated for adipose neurovascular labeling. For complete details on the use and execution of this protocol, please refer to Willows et al. (2021).


MATERIALS AND EQUIPMENT
1. You will need access to a cold room (4 C) and an orbital shaker. 2. It is recommended to have an aspirator setup to remove buffers from wells between washes/incubations. 3. A pair of forceps and dissecting scissors. 4. Access to an epifluorescence and/or confocal microscope. In this step you will remove intact axillary and inguinal subcutaneous white adipose tissues from mice for immunostaining. For a demonstration of an axillary and inguinal subcutaneous white adipose tissue (scWAT) dissection see Methods video S1.
Note: This protocol is performed without perfusion. Any potential benefits or drawbacks of perfusion with this protocol have not been tested thoroughly at this time.
CRITICAL: Maintain inguinal scWAT orientation for all future steps, up to and including mounting. Most importantly, make note of whether the tissue was taken from mouseleft or mouse-right as the tissue will always curve towards the body when placed medial side up (congruent to tissue positioning during dissection, Figure 1). Additionally, the axillary end of the tissue will be wider than the posterior end. With these notes in mind, if it is known which side of the mouse the depot was excised from, it can always be returned to appropriate orientation when needed.
CRITICAL: Maintain axillary scWAT orientation for all future steps, up to and including mounting. The Axillary depot is more difficult to orientate than the inguinal depot as there are no obvious features for orientation. The most important step is to excise the tissue and place it medial side up into a well noting mouse-left or mouse-right. The axillary end of each tissue will have characteristic browning (in most cases) which can help orientate further, but this will be obscured with the addition of autofluorescence quenching stains described later.
2. Excise inguinal scWAT depot. a. Place mouse ventral side up, spray with 70% EtOH. b. Make a central incision from the base of the tail extending to the top of the sternum, being sure not to penetrate the underlying peritoneal cavity. c. Gently use forceps and thumbs to pull ventral skin away from both flanks, pulling slowly with even pressure, making sure not to damage the underlying adipose tissue. The skin can be pinned back (as demonstrated in Methods video S1) to better expose the adipose depots. d. Identify the inguinal scWAT depot against the skin besides each hip (Figure 1 -boundaries of each depot). Use scissors to cut the large vasculature leading to the depot and then trace the edges of the tissue, cutting along the way. e. Use forceps to gently roll the adipose away from the skin -cutting the anchoring connective tissue as you go. Continue until the entire depot is free from the skin. f. Weigh the depot and place it into a 6-well plate with 3-5 mL ice-cold 2% PFA and fix at 4 C. Fix the tissue without agitation to prevent the tissue from folding onto itself during the fixation process. i. %0.08g ; fix for 8 h.
Note: Once tissues are adequately fixed without agitation, they are unlikely to fold during incubations performed on an orbital shaker as utilized in later steps. 3. Excise axillary scWAT depot. a. Identify axillary depot underneath both forelimbs ( Figure 1). b. Use forceps to pull away a thin muscle layer enveloping the dorsolateral portion of each axillary depot. c. Use scissors to trace the edges of the tissue, cutting along the way. d. Use forceps to gently roll the adipose tissue away from the skin -cutting the anchoring connective tissue as you go. Continue until the entire depot is free from the skin. e. Weigh the depot and place it into a 6-well plate with 3-5 mL ice-cold 2% PFA to fix at 4 C. Fix the tissue without agitation. i. %0.08g; fix for 8 h.

Reducing tissue thickness and autofluorescence
Timing: Z-depth reduction, 1 h 30 min; permeabilization, 12 h; autofluorescence quenching, 4 h-1 day Adipose tissue is notoriously difficult to image due to its size and inherent autofluorescence. In this step we will be using a combination of Z-depth reduction and various autofluorescence quenching techniques to greatly reduce tissue background autofluorescence. It should be noted that hypertrophic adipocytes, and human adipocytes, may be more prone to cell lysis on the surface of the tissue with this technique. In some instances, whole tissue imaging is not required, and instead smaller Figure 1. Excising intact subcutaneous white adipose tissue (scWAT) depots from mice A male C57BL/6J mouse had skin pinned back to display inguinal and axillary scWAT depots before and after excision. White dashed lines are used to accentuate tissue boundaries. Tissues were kept medial side up (as in the intact scWAT image) and directionality was maintained; anterior (A, towards head) and posterior (P, towards tail).

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representative tissue pieces can be used. We have also outlined this alternative approach as an optional step below.
Note: All washes and incubations are performed on an orbital shaker.
a. Following fixation, aspirate PFA from each well of the 6-well plate and add 13 PBS w/heparin. Wash tissues for an hour at 4 C, replacing 13 PBS w/heparin after 30 min. Heparin is used here to help remove highly autofluorescent blood cells from the vasculature.
Pause point: The conclusion of any series of 13 PBS washes throughout this protocol marks a good pause point if desired (e.g. washes following fixation, autofluorescence quenching, or antibody staining.) Tissues can remain in 13 PBS for several days with no observed detriment to staining outcome. It is prudent to do antibody incubations immediately prior to imaging, for best results.
b. Use forceps to take the tissue out of PBS and lay it flat onto the center of a large (75 3 51 mm) glass slide. c. Use a transfer pipette to add 2-3 drops of PBS onto the tissue to prevent the tissue from drying out. d. Take a second large glass slide and place it on top of the tissue; sandwiching the tissue betwixt. e. Take two large (5 cm wide) binder clips and clamp them onto the slides ( Figure 2). f. Allow tissues to remain compressed for 1 h 30 min at 4 C. g. Remove binder clips and gently separate the two slides without damaging the tissue. See Optional: To entirely avoid compressing tissues, Z-depth reduction can be removed from this protocol by cutting each adipose tissue into 4-5 smaller pieces and following through with the remaining steps until mounting. Mount each piece of tissue onto a concave slide (0.8-1.9 mm deep, depending on tissue thickness). We do not recommend this alternative approach unless it is determined that tissue compression must be completely avoided. Only the most superficial structures will be visible. See Willows et al. (2021) for a comparison of both appropaches.
5. Blocking and Permeabilization. a. Once the slides are separated place the tissue back into a 6-well plate with 3-5 mL blocking buffer at 4 C for 16-24 h to reduce non-specific binding of antibodies.
Note: Here, bovine serum albumin is used as the blocking agent with great effect. Alternatively, using secondary-matched sera as a blocking agent is common with many other immunostaining protocols, but has not yet been thoroughly evaluated for use with this protocol. i. Pro -reduces autofluorescence without affecting fluorophore intensity. ii. Con -de/rehydration steps add several hours, and bleaching adds an additional day to the protocol. MeOH preparation has been demonstrated not to be compatible with some antibodies as it can denature epitopes (Renier et al., 2014) and should be tested and validated on a case by case basis. e. Option 4 -Room Temperature: doing all future steps at $20 C (even when recommended at 4 C) will reduce tissue autofluorescence, but only slightly. i. Pro -shortens protocol by at least 4 h while still offering some level of autofluorescence reduction. ii. Con -the least effective means of reducing autofluorescence and performing all future steps at $20 C can result in increased non-specific binding of antibodies.
Optional: Washes following autofluorescence quenching techniques can be performed in 13 PBS w/ heparin to further reduce vascular autofluorescence. Additionally, sodium azide can be added to PBS at a concentration of 0.02% for all future washes to prevent microbial growth.

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CRITICAL: The addition of non-ionic detergents (Triton X-100, Tween 20) into wash buffers is common to many immunostaining protocols, but if using typogen black or TrueBlack autofluorescence quenching, avoid the addition of these detergents as they can wash out these stains from the tissue, greatly reducing their quenching potential. The use of detergents has not been tested with the other quenching techniques.

Fluorescent labeling
Timing: 1-7 days The following steps outline how to fluorescently label Z-depth reduced adipose tissues or tissue blocks with primary antibodies, secondary antibodies, isolectin-IB 4 , and DAPI. It is important that, to reduce non-specific binding and cross reactivity, co-staining follows the order outlined below.
Note: All washes and incubations are performed on an orbital shaker.
7. Using a Direct Reporter with Endogenous Fluorescence a. For mouse reporters with high signal intensity you can proceed directly to step 12 or if co-labeling is desired, proceed to step 8. b. For mouse reporters with low signal intensity it is recommended to boost the endogenous signal with a fluorophore conjugated antibody raised against the fluorescent protein of interest (i.e., anti-GFP conjugated Alexa488). This should be conducted immediately after step 9 is completed. 8. Primary Antibodies. a. To make primary antibody solution, you will dilute desired primary antibody in 13 PBS. Dilutions are antibody specific but typically range from 1:100 to 1:500. For antibodies not validated in this protocol, we recommend performing an antibody titration to find what works best. When titrating antibodies for this protocol, it is best to start with what works well for thin sections as this can provide decent enough staining. However, concentrating this further by two-fold typically results in the best outcomes. Concentrations for antibodies that we have validated with this protocol can be found in the key resources table.
CRITICAL: Non-ionic detergents are commonly added to antibody solutions to increase antibody permeation and reduce non-specific binding, however, as mentioned previously, detergents can reduce the effectiveness of typogen black and TrueBlack, and has not been tested with the other quenching techniques.
b. You will need 3-5 mL of antibody solution per tissue, 3 mL being the absolute minimum if using a 6-well plate (see ''troubleshooting''.) c. Incubate tissues in primary antibody solution for 2 days at 4 C. d. After incubation, wash tissues in 13 PBS for 4 h at 4 C, replacing PBS every 1 h.
Note: Perform sequential staining of antibodies when co-staining.
Note: Only validated antibodies, tested in a Western Blot for specificity, and ideally with knock-out tissues and proper tissue controls, should be used for this procedure.
9. Secondary Antibodies a. If not using a directly conjugated primary antibody, dilute the desired secondary antibody in 13 PBS. Again, dilutions will be antibody and fluorophore specific, and should be optimized by titration. Secondary antibody concentrations tend to be identical to that used for immunostaining thin sections as recommended by the vendor. Only Alexa Fluor conjugated antibodies have been validated for this protocol. See validated secondary antibody concentrations in the key resources table. b. Incubate tissues in secondary antibody solution for 16-24 h at 4 C. c. After, wash tissues in 13 PBS for 4 h at 4 C, replacing PBS every 1 h. d. If co-staining with additional primary antibodies go back to step 8. If not, continue to step 10. CRITICAL: We recommend avoiding the use of anti-mouse secondary antibodies. In most cases, non-specific binding has been unavoidable even with mouse-on-mouse blocking solutions. We recommend using antibodies raised in mice that are directly conjugated to a fluorophore, or preferably, using primary antibodies raised in alternative species such as rabbits. a. If nuclear labeling is desired, start by making DAPI working solution from stock (described in materials and equipment), to be used fresh. b. Incubate tissues in 3-5 mL DAPI working solution for 1 h at $20 C. c. Wash in 13 PBS for 2 h at $20 C, replacing PBS after 1 h.

Mounting tissues
Timing: 3 days A benefit of our protocol is the ability to permanently mount entire subcutaneous adipose tissues onto glass slides. Below we outline the steps for mounting.
a. Label the top of a large glass slide with mouse and tissue identifying information. b. Remove tissue from 6-well plate and place it at the center of the large glass slide. c. Add a generous amount (4-6 drops) of glycerol based mounting fluid to the tissue and place a large (48 3 60 mm) glass coverslip over the tissue.
Note: Glycerol-based mountants are more compatible with lipid laden tissue, and the added viscosity aids in adhering the coverslip to the slide. Incomplete coverslip adherence will result in tissues drying out, tissue oxidation, and decreased fluorophore lifespan.
d. Apply gentle pressure to the coverslip to help it adhere to the tissue and slide. This is required for the next step to prevent the coverslip from sliding off the tissue. e. Place the coverslipped slides underneath uniform weight (15-20 kg) for 3 days. Ten textbooks placed over a series of slides works very well. f. After 3 days, remove the slide from underneath the textbooks and seal the edges with nail polish (Figure 4). g. Let dry for 5 min and proceed to imaging.
Note: For best outcome we recommend imaging within a week after mounting is complete. However, tissues are typically viable for imaging up to several weeks after mounting, and in some cases even several months.

Imaging
Timing: 1 min-40 h Unique to this tissue processing approach is the ability to image whole adipose tissues effectively on standard widefield epifluorescence microscopes as well as confocal microscopes. Below are descriptions for best utilizing both methods of imaging.
13. Widefield Epifluorescence Microscopy a. Application: i. Suitable for single channel imaging, most qualitative analysis, and answering 'yes or no' questions through representative images. Fast time to image with only minutes needed to set up and capture several images. b. Magnifications: i. Images of adipose tissue can be captured on widefield microscopes easily using up to 403 objectives. Higher magnifications are possible but the associated decrease in working distance will make it a challenge to capture an in-focus image of large tissues. i. Even with autofluorescence quenching, background will still be present on epifluorescence microscopes (see ''troubleshooting''). Strong staining and bright fluorophores can help overcome this, but confocal microscopy will further help to reduce background. 14. Confocal Microscopy a. Application: i. Suitable for quantification, demonstrating colocalization, and generating 3D images of specific regions or entire tissues to capture total neurovascular structure. b. Magnifications: i. Confocal microscopy is capable of imaging whole adipose tissues with up to 633 objectives without difficulty -further increased with confocal zoom. We have not tried higher objective magnifications at this time. c. Tiled Z-stacks: i. Entire adipose depots can be imaged in a series of Z-stacks extending through the entire tissue thickness. These can be tiled together to create one large 3D image. These can then be maximum intensity projected to create a 2D representation of the 3D data. These tiled z-stacks of whole tissues can be very time consuming to capture; taking anywhere from 8 to 40 h depending on the size of the tissue, number of fluorophores being captured, and the desired XY and Z resolutions.

EXPECTED OUTCOMES
With this protocol you will be able to fix and immunostain entire subcutaneous adipose depots and effectively image them without optical clearing on widefield and confocal microscopes. Below we

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have provided examples of images captured using this protocol ( Figure 5). For additional examples, see previous publications using this protocol (Blaszkiewicz et al., 2019(Blaszkiewicz et al., , 2020Willows et al., 2021).

QUANTIFICATION AND STATISTICAL ANALYSIS
Additionally, we have developed a means to quantify whole tissue innervation that is complementary to the tissue processing and imaging techniques described here. For further details see Willows et al. (2021) and access the code at https://github.com/ktownsendlab/willows_et_al-2020.

LIMITATIONS
This protocol was developed with adipose nerve and neurovascular immunostaining in mind. We strongly believe that it is applicable for any number of research goals but will need to be validated  for uses outside of the scope of this protocol. The Z-depth reduction approach can potentially alter tissue morphology, so we recommend researchers implementing this technique conduct their own observational analysis to ensure that Z-depth reduction does not create artefact in the structures they analyze. Z-depth reduction, by definition, compresses tissue in the Z-axis and results in reduced Z-axis spatial resolution. (Willows et al., 2021) provides comparisons of adipocyte size with and without this technique, for example.

TROUBLESHOOTING
Problem 1 Autofluorescence quenching with typogen black or TrueBlack reduces signal intensity and hides fluorescing structures.

Potential solution
Try using more dilute concentrations of these quenching agents as they can mask fluorescence signal. We have demonstrated the use of a few different concentrations (Willows et al., 2021). Alternatively, using hydrogen peroxide bleaching or negating autofluorescence quenching entirely may be preferable in certain circumstances.

Problem 2
Tissues don't compress sufficiently when coverslip is added during mounting.

Potential solution
Reduce fixation time. Overly fixed tissues will be a bit more rigid and tend to 'slide around' when gentle pressure is applied with the coverslip.

Problem 3
High background autofluorescence even after quenching.

Potential solution
The use of very bright fluorophores at high concentrations (anti-rabbit IgG Alex Fluor 647 Plus, 1:200) will overcome most background signal if imaged on a confocal microscope.

Problem 4
Low intensity fluorescence signal.

Potential solution
Increase the volume of antibody solutions to 5 mL and avoid conjugated primary antibodies if possible. Indirectly labeling with secondary antibodies will provide greater signal intensity. Additionally, increasing incubation times may also help.

Problem 5
Imaging entire whole depot innervation is taking 'too long.'

Potential solution
Adjust image acquisition settings. Optimizing image capture settings for a tiled Z max projection of an entire tissue can greatly reduce acquisition time. Settings will vary between confocal systems, but here are what we recommend for a Leica SP8 and Leica Stellaris 5 systems: Resolution 720 3 720 (can be reduced further if needed), scan speed 600 Hz, pinhole 1AU, scanned bi-directionally, 2line average, Z-step size of 10-16 mm, tiled with a 10% tile overlap. Increase laser intensity and gain as needed. DAPI and Alexa647 can be imaged simultaneously, but all other fluorophores must be imaged sequentially by line (or by frame if using DAPI; long depletion time can cause bleed through.) Using a microscope that can 'lasso' areas of interest for tiling instead of imaging entire rectangles will prevent wasting hours on imaging empty space.