To visualize macromolecule synthesis, we injected click-it compatible monomer analogs (Table 1) intraperitonially 3 hr before sample collection. During this time, analogs metabolically incorporated into nascently synthesized macromolecules, resulting in in vivo labeling of macromolecule synthesis. Our group has found that collection of tissue after 3 hr provides sufficiently strong metabolic labeling of DNA synthesis (EdU), transcription (EU), translation (AHA), and protein glycosylation (GlcNAz). These labeled macromolecules contain either azide- or alkyne-modified monomers that can be detected with click-it compatible fluorescent molecules, enabling imaging of nascent macromolecules in whole-mount tissues with LSFM (Figure 1B–C). We show that our whole-mount method can be used at the organismal level to visualize the whole torso of a stage 52 axolotl larvae, where we observe proliferating cells in the developing limbs and hematopoiesis within the spleen (Figure 1B). To demonstrate the multiplexing capabilities of the approach, modified monomer analogs with disparate functional groups (EdU/AHA, 5-EU/AHA, EdU/GlcNAz, 5-EU/GlcNAz) were co-injected and visualized with LSFM (Figure 2A–D, Videos 1–3. For color blind accessible images, see Figure 2—figure supplement 1). whole-mount samples were comparable to 2D longitudinal tissue sections of the same stains (Figure 2I–L), showing that our method generates similar results in both whole-mount and tissue sections (Figure 2—figure supplement 2) with subcellular resolution (Figure 2—figure supplement 3). Furthermore, the number of EdU⁺ cells per blastema area in a single Z slice obtained by LSFM (EdU-AHA: 1954 cells/mm², EdU-GlcNAz: 1458 cells/mm²) is similar to that of a tissue section obtained by confocal microscopy (EdU-AHA: 1272 cells/mm², EdU-GlcNAz: 1349 cells/mm²). We finally demonstrate the specificity of GlcNAz incorporation by pretreating a GlcNAz-specific antibody on tissue sections collected from GlcNAz injected animals. By doing so, the subsequent click-it reaction was prevented due to the antibodies likely sterically hindering alkyne fluorophores from accessing the azide functional groups (Figure 2—figure supplement 4).

An advantage of our method is that staining whole-mount tissues eliminates the need for sectioning, reducing the potential inconsistencies that arise as a result of the sectioning process (uneven tissue, different cutting planes, etc.). Additionally, traditional methods of obtaining 3D images of thick tissues with confocal microscopy are impractically slow when imaging hundreds of images in a single stack. LSFM allows for more rapid imaging of whole samples, requiring only minutes to image each sample. However, several considerations exist when imaging whole tissue in 3D with LSFM. Stain penetrates slower in whole 3D tissues compared to 10–20 µm thick tissue sections, requiring longer staining times. An advantage of our method is that the click reaction requires molecularly diminutive reagents that readily pass through cell and nuclear membranes, ensuring stain penetration in the center of dense tissues. Different refractive indices between disparate cellular components and the imaging media cause light to scatter, which can reduce the resolution and brightness of 3D images (Fadero and Maddox, 2017). To improve image resolution and light penetration, overnight refractive index matching with 67% 2,2’-thiodiethanol (TDE) can sufficiently clear axolotl limbs for imaging with LSFM. TDE was chosen due to its simplicity, cost, safety, and compatibility with common imaging modalities. This clearing method rapidly and effectively improves the signal-to-noise ratio of stained samples compared to imaging in PBS (Figure 2—figure supplement 5). Tissue morphology was minimally disturbed after clearing, with only mild shrinkage of the blastema epithelium observed. With careful attention to these challenges, our method provides a means to obtain high-quality 3D images from tissues 1 mm in depth in less than 10 min per sample with clear, consistent staining (Figure 2A–H, Figure 2—figure supplement 3).

To demonstrate that our whole-mount click-it method can obtain quantifiable data on the organ, cellular, and molecular levels of organization, we applied our technique to the regenerating humerus. After regenerating for 35 d, axolotls with mid-humeral amputations were injected with EdU/AHA to identify cells within the humerus undergoing DNA synthesis (EdU) and protein translation (AHA). We observed EdU staining in chondrocytes distal to the amputation plane and AHA staining in the humerus perichondrium (Figure 3A). We outline a multiscale, quantitative pipeline that leverages the staining patterns of these macromolecules for analysis of 3D humerus morphology and 3D macromolecule synthesis. This workflow combines available plugins in Fiji (Schindelin et al., 2012) and scripts developed in Fiji and Matlab (The MathWorks, 2019) for the data analysis process (see supplementary information for a detailed description).

For 3D organ level analysis of macromolecule synthesis, AHA staining provided an adequate outline of the humerus, allowing us to segment its 3D morphology (Figure 3A–B). We quantified organ shape by assessing cross-sectional area (Figure 4A’) and circularity (Figure 4A”) of the segmented humerus along the proximodistal axis with BoneJ (Doube et al., 2010; Figure 4A). These measurements of 3D organ size can be obtained with other methods such as microCT and focused ion beam scanning electron microscopy (FIB-SEM). However, microCT is unable to image every tissue due to stain limitations and uses hazardous radiation, while FIB-SEM can capture 3D surface topography but not the entire organ morphology. Our method has the capability to fully image the 3D structure of entire organs (size permitting) without the need for radiation while simultaneously capturing information from both the cellular and molecular levels of organization.

On the cellular level, we segmented the 3D morphology of proliferating chondrocytes based on EdU staining using the Trainable Weka Segmentation 3D plugin (Arganda-Carreras et al., 2017; Figure 4B”). With this segmentation, we identify highly condensed regions of cells undergoing rapid rates of DNA synthesis (Figure 4B). From these data, we observe cells synthesizing DNA most abundantly distal to the plane of amputation, as expected for dividing chondrocytes. Traditionally, cell quantification as such is conducted on 2D tissue sections. In heterogenous tissues, however, cells are distributed non-uniformly; 2D sampling may not accurately capture the cellular distribution and cannot be used to determine cell volume or shape. Our whole-mount staining method allows quantification of cells within an entire 3D tissue, resulting in a more accurate assessment of cell density within heterogenous tissues.

To demonstrate the quantitative molecular analysis of 3D macromolecule synthesis, we assessed the staining intensity of EdU in slices along the proximodistal axis of the regenerating humerus (Figure 4C). EdU intensity represents the rate at which a cell undergoes DNA synthesis, which is one of the first steps of cell division and proliferation. Proliferating cells rapidly synthesize DNA, which enables these cells to integrate EdU into nascent DNA strands. This provides ample opportunities for covalent linkage of fluorescent molecules to the DNA strand. Thus, higher pixel values within EdU⁺ cells are representative of more DNA synthesis in the cell. These data show that EdU intensity is strongest in the regions distal to the amputation plane of the humerus (Figure 4C’), providing a quantitative measure to assess macromolecule synthesis on the molecular level. Within a tissue, cells synthesize macromolecules heterogeneously, reflected by different fluorescence values between cells. Thus, quantifying macromolecule synthesis based on the presence/absence of signal instead of fluorescence does not account for variability in macromolecule synthesis rates among cells. To compound this issue, quantifying fluorescence in tissue sections only represents the rate of macromolecule synthesis from a fraction of cells in larger, heterogenous tissues. Our method provides a means to capture this molecular heterogeneity in 3D samples, allowing us to observe whole 3D regions in the regenerating humerus that synthesize DNA more rapidly than others, which further demonstrates the utility of our whole-mount click staining method.

Taken together, these results demonstrate that our method can provide quantifiable data on the organ, cellular, and molecular levels of organization. This highlights the novelty of our method, as we have not found previous examples of multiscale analysis as outlined here. We foresee this multiscale, quantitative analysis having broad applications in the examination of dynamic cell processes in 3D, such as in cancer metabolism and mammalian neurogenesis or other fields where macromolecule synthesis is traditionally studied in tissue sections.

To demonstrate that our method is sensitive enough to detect subtle changes in macromolecule synthesis in vivo, we quantified the difference in EdU intensity between limbs regenerating with and without a nerve supply. Blastema cells are thought to have a cell cycle length of 40–50 hr, with S phase approximately 30 hr (Tassava et al., 1987). In newts, it has been shown that amino acid, RNA, and DNA analog incorporation decreases approximately 30% by 24 hr after denervation (Singer and Caston, 1972). Based upon these estimates, we amputated both forelimbs at the mid-humerus and denervated the left limb at the brachial plexus 24 hr before collection. This timepoint for denervation was chosen because it should be sufficiently long to have an impact on DNA, RNA, and protein synthesis rates. At 6, 9, 12, 15, 18, and 25 d post-amputation (dpa) animals were pulsed with EdU for 3 hr before collection to label proliferating blastema cells (Figure 5A). We chose 3 hr of EdU incorporation to provide a snapshot of DNA synthesis in the 30 hr total S phase of blastema cells. LSFM was used to image samples (Figure 5B), ensuring pixel resolution was consistent between samples. We quantified DNA synthesis in denervated limbs compared to innervated limbs by creating a 175 × 175 × 175 µm cube 250 µm from the distal most tip of the blastema (Figure 6A). The size and location of the quantification cube was chosen to maximize the number of EdU⁺ mesenchymal cells and to exclude any epithelial cells. Although we limited the cube to the size of the smallest blastema, these parameters can be customized depending on the size of the blastemas. For this study, we estimate 50–150 blastema cells are found within the cube. From these results, we observed a marked decrease in blastema EdU incorporation due to denervation at 9, 12, and 15dpa (Figure 6B, Figure 6—figure supplement 1), demonstrating that our whole-mount staining approach is capable of detecting changes in macromolecule synthesis after biological perturbations. One potential limitation of our method in the blastema is the inability to perform single cell segmentation. This is due to the density and abundance of cells within the blastema. We predict that higher magnification imaging and deep learning segmentation techniques may overcome this limitation but will significantly increase imaging time and file size. For comparison, we have included estimates of imaging time and file sizes for confocal microscopy with a 20X objective and LSFM with a 5× objective (Supplementary file 1).

Axolotls were either bred in captivity at Northeastern University or purchased from the Ambystoma Genetic Stock Center at the University of Kentucky. Experiments were performed in accordance with the Northeastern University Institutional Animal Care and Use Committee. Animals were grown to 4–6 cm (Mean 5.3 cm, SD 0.36) and 1–1.5 g (Mean 1.3 g, SD 0.19 g) for use in all studies. For all experiments, animals were anesthetized by treatment of 0.01% benzocaine until visually immobilized. Limbs were amputated either at the distal end of the zeugopod or midway through the stylopod, and bones were trimmed below the amputation plane to allow for uniform growth. At the date of collection, animals were reanesthetized and injected with either 5-ethynyl-2′-deoxyuridine (EdU) to identify proliferating cells (8.0 µg/g animal), 5-Ethynyl Uridine (5-EU) to label RNA (270.0 µg/g animal), L-Azidohomoalanine (AHA) to label protein (180.59 µg/g animal), or N-azidoacetylglucosamine-tetraacylated (GlcNAz) to label glycosylated proteins (430 µg/g animal) alone or simultaneously in the following combinations: EdU/AHA, EdU/GlcNAz, 5-EU/AHA, 5-EU/GlcNAz (Table 1). All monomer analogs were purchased from www.clickchemistrytools.com and resuspended in DMSO at the following concentrations: EdU- 300 mM, 5-EU- 100 mM, AHA- 100 mM, GlcNAz- 100 mM. Stocks were further diluted in 1× phosphate buffered saline (PBS) for injection. After 3hr of analog incorporation, limbs were collected from the upper stylopod and fixed in 4% paraformaldehyde (PFA) (diluted in 1× PBS) at 4°C overnight. If limbs were denervated, the nerve supply was severed at the brachial plexus 24 hr before tissue collection.

Following fixation in 4% PFA, samples were washed three times with 1X PBS at room temperature (~23°C) for 5 min. Samples were dehydrated in an increasing methanol series at room temperature starting with 25% methanol (diluted in 1× PBS), 50% methanol, 75% methanol, and 100% methanol for 5 min at each step. Samples could then be stored in 100% methanol indefinitely at −20°C. For staining, samples were rehydrated in a decreasing methanol series starting with 75% methanol (diluted in 1× PBS), 50% methanol, 25% methanol, and finally placed in 100% 1× PBS for 5 min at each step. Samples were then washed three times with 1× PBST (1× PBS with 0.1% Triton) for 5 min at room temperature. To aid in clearing, samples were washed in 0.5% trypsin (diluted in 1× PBS) for 30–90 min on a rocker at room temperature, or until the sample appeared translucent. Samples were washed three times at room temperature for 5 min with deionized water, then washed in 100% acetone for 20 min at −20°C and washed with deionized water again for 10 min. Samples were washed in 1× PBST three times at room temperature for 5 min before applying click-it cocktail for overnight at room temperature. The click-it cocktail was made in 500 µl of 1× TRIS buffered saline as follows: 50 µL 1M sodium ascorbate (100 mM final), 20 µL 100 mM CuSO₄ (4 mM final), and 2 µL 500 µM azide- or alkyne-modified Alexa Flour (2 µM final), combined in order as listed. After the first round of staining, samples were washed at room temperature six times for 30 min with rocking. For double-labelling, samples were again placed in the click-it cocktail with a different fluorescent dye to stain for the second analog at room temperature overnight. Both rounds of staining were conducted in the dark to prevent photodegradation of fluorescent molecules. For staining with DAPI, samples were washed three times for 5 min with 1× PBS, then placed in 2.86 µM DAPI for 4 d at room temperature. Samples were washed three times for 20 min with 1× PBS and left in 1× PBS at 4°C for short-term storage before imaging with LSFM. whole-mount samples were cleared with 67% TDE (diluted in 1× PBS) overnight at room temperature in the dark.

Following fixation in 4% PFA, samples were washed three times in 1× PBS each for 5 min, and cryoprotected in 30% sucrose on a rocker until the tissue fully sank. Samples were removed form sucrose and briefly washed in optimal cutting temperature (OCT) compound before mounting in OCT compound and frozen at −80°C. A cryostat was used to obtain 10 µm sections, and slides were baked at 65°C for 15 min. Slides were then washed with water for 30 min at room temperature to remove residual OCT. Slides were washed once with 1× PBS for 5 min at room temperature. The click-it cocktail (same as above) was applied to the slides and incubated at room temperature in the dark for 30 min. If staining for a second macromolecule, slides were washed five times for 5 min with 1× PBS at room temperature. The samples were then stained for 30 min at room temperature in the dark using the above click-it cocktail with a different a fluorophore dye. Following the final click-it reaction, slides were washed once with 1× PBS at room temperature for 5 min, then stained with 2.86 µM DAPI for 5 min at room temperature. Slides were washed again with 1× PBS for 5 min at room temperature and water for 5 min at room temperature and mounted with SlowFade Gold Antifade Mountant. Slides were imaged using a Zeiss LSM800 confocal microscope.

All 3D images were acquired using a Zeiss light sheet Z.1 microscope paired with Zen software. Unless otherwise indicated, samples were cleared and imaged in 67% TDE. Post-processing for visualization purposes was performed with Arivis Vision4D v3.1.4 on a workstation with a 64-bit Windows Embedded Standard operating system, and an Intel(R) Xeon(R) CPU E5-2620 v3 @ 2.40 GhZ (two processors), 128 GB RAM, and NVIDIA Quadro K2200 GPU. Sub-volumes were stitched together with the Tile Sorter in Arivis, using the manual projection option. Volume fusion was performed through automatic landmark registration of manually selected points for alignment. For visualization, background intensity was corrected using the automatic functionality.

All data were processed on desktop computers with the aid of Fiji (Schindelin et al., 2012) and Matlab (The MathWorks, 2019). The custom Fiji scripts and Matlab codes used are available in the supplementary material. We performed all analyses on unprocessed. czi files acquired directly from the light sheet microscope.