We thank all members of the BSU staff at the Kennedy Institute of Rheumatology, particularly Albertino Bonifacio, for animal husbandry. We thank the Oxford-ZEISS Centre of Excellence of Biomedical Imaging for assistance in using the microscope, in particular, Dr. Jacky (Ka Long) Ko for help in developing the image analysis pipeline. This work was supported by the Kennedy Trust of Rheumatology Research studentship (Johnson) and the Biotechnology and Biological Sciences Research Council (BBSRC, BB/X007049/1).

Animal husbandry and experiments were conducted in accordance with the University of Oxford ethical frameworks and under a Project license as granted by the UK Home Office. This protocol was performed on mice between the ages of 2 and 10 weeks but would work in the cartilage of mice beyond this age.
1. Tissue collection
<ol>
	<li>Euthanize mice by filling a box containing the mice with carbon dioxide up to 80% by volume and maintaining its concentration for 5 min. Perform cervical dislocation.</li>
	<li>Dissect out the tissue of interest. 
	NOTE: Below is a protocol for dissecting the hind leg knees, including the tibial and femoral growth plates and articular cartilage.
	<ol>
		<li>Incise the skin at the foot and cut to the top of the thigh. Remove skin from the incision line with forceps.</li>
		<li>Trim the muscle and fat tissue from the legs.</li>
		<li>To detach the leg from the body, cut cleanly through the femur at the top of the hip. Cut the foot at the bottom of the tibia.</li>
		<li>Remove any excessive muscle and fat tissue left on the leg.</li>
	</ol>
	</li>
</ol>
2. Tissue fixation and embedding
NOTE: Throughout the protocol, take care to keep the tissue in the dark to avoid fluorescent signal loss.
<ol>
	<li>Fix the tissue in 10% formalin at room temperature for 4 h.</li>
	<li>Transfer the tissues to 2.5% formalin at 4 &#176;C overnight.</li>
	<li>Transfer the tissues to 30% sucrose in distilled water (dH2O), and gently rotate at room temperature overnight or for 3 days at 4 &#176;C. Fill the tubes to the top to ensure complete immersion of the tissue during the rotation. Following this incubation, the tissue will sink to the bottom of the tube.</li>
	<li>Place a small amount of dry ice in a separate box and position the cryomold on top. Trace a line of super cryoembedding medium (SCEM) and place the limb along it, with the kneecap facing down and opening the tibia and femur apart for the leg to be as flat as possible. Hold in place with forceps until the SCEM has started to solidify and the limb no longer moves when not held down.</li>
	<li>Cover completely with SCEM, filling the mold to the top. Place the sample in the dry ice box, keeping the sample flat to avoid movement of the tissue, until the SCEM completely solidifies.</li>
	<li>Transfer to -20 &#176;C.</li>
</ol>
3. Sectioning
<ol>
	<li>Remove the block from the mold and trim to a size that fits on the chuck of a cryostat. Attach the block to the chuck by applying SCEM on the chuck and placing the block on top.</li>
	<li>Allow to solidify for at least 5 min at -20 &#176;C in the cryostat.</li>
	<li>Attach the chuck to the sample holder and a blade to the blade holder.</li>
	<li>Trim the block until the tibial growth plate is reached. Ideally, section with the tibia pointing upwards.</li>
	<li>Cut cryofilm tape to a size that fits the sample with enough space surrounding it. Mount the cryofilm to the cut surface, ensuring tight adhesion of the cryofilm with a fitting tool. Cut 40-60 &#181;m-thick sections to obtain two-cell thick sections. Collect the tape using forceps and place on a slide, sample side up, and the tibial growth plate in line with the short slide of the slide.</li>
	<li>Store the sample slides at -20 &#176;C.</li>
</ol>
4. Slide preparation and mounting
<ol>
	<li>Thaw the slide horizontally at room temperature.</li>
	<li>Wash the slide with a drop of PBS for 3 x 5 min to remove the SCEM.</li>
	<li>Stain with 10 &#181;M DAPI for 1 min.</li>
	<li>Wash the slide with a drop of PBS for 2 x 5 min.</li>
	<li>Mount, cover with a coverslip, and leave at room temperature overnight. Seal the edges of the coverslip with coverslip sealant.</li>
	<li>Image the following day.</li>
</ol>
5. Imaging by confocal microscopy
NOTE: The resolution of the 40x/1.4 objective on the confocal microscope used is 240 nm.
<ol>
	<li>Set up the microscope.
	<ol>
		<li>Turn on the microscope and open the software. Click on the System button.</li>
		<li>Under the Acquisition tab, click on Smart Setup to set up the channels. Select and add the GFP, mCherry, and DAPI channels. Select the Airyscan option and the SR8Y function in the Airyscan triangle (between resolution and speed-multiplex 8Y). Click on Best Signal | OK (Supplemental Figure S1A).</li>
		<li>In the Locate tab, under Microscope Control, select the 40x/1.4 objective with oil immersion.</li>
	</ol>
	</li>
	<li>Locate the region of interest.
	<ol>
		<li>Place the slide in the slide holder and bring the objective up until the oil touches the coverslip.</li>
		<li>Position the growth plate or other region of interest underneath the light path.</li>
		<li>In the Locate tab, click on Fluorescence, select DAPI,&#160;and using the eyepiece, move the slide holder with the joystick to position the growth plate or region of interest under the field of view.</li>
		<li>In the Acquisition tab, start by selecting the DAPI channel only.</li>
		<li>Click on Continuous to view the image of that channel on the screen and ensure the growth plate is in view.</li>
		<li>Open the Tile viewer by clicking on Show viewer in the Tiles tab. Add the number of tiles required (usually 1 tile in the x direction and 3-5 in the y direction) and click on Preview and Start (Supplemental Figure S1B).</li>
		<li>Check that the tiles cover the three zones of the growth plate-resting, proliferation, and hypertrophic. Adjust the tile position and number as required.</li>
	</ol>
	</li>
	<li>Set up the imaging parameters.
	<ol>
		<li>Click on Continuous in the Acquisition tab. In the Channels tab, adjust the Gain (Master) and Laser power to view the nuclei clearly on the screen. Allow the Airyscan Detector to align. Open the Airyscan Detector Adjustment window by clicking on the tiled rosette symbol at the bottom of the window; once it has aligned, all the tiles will turn green (Supplemental Figure S1C).</li>
		<li>Select the GFP and mCherry channels individually and repeat these steps-setting the Gain (Master), Laser power, and align the Airyscan Detector.</li>
		<li>Select all three channels and click on Continuous. Allow the Airyscan Detector to align.</li>
		<li>In the Acquisition Mode tab, define the Image Size, Pixel, Size, Frame Size, Speed, and Averaging values. See Supplemental Figure S1D,E for values of Acquisition Mode settings, Gain (Master), and Laser power for the channels used.</li>
	</ol>
	</li>
	<li>Acquire images.
	<ol>
		<li>In the Acquisition tab, click on the Z-Stack tick-box.</li>
		<li>With just the DAPI channel selected, click on Continuous to view the image on the screen. Move the objective down to the closest plane to be imaged. In the Z-Stack tab, click on Set First plane, focus away to the last plane to be viewed, and click on Set Last plane. Image 30-40 &#181;m z-stacks.</li>
		<li>Set the interval to 0.15 &#181;m.</li>
		<li>Move to the center of the z-stack. In the Tiles tab, under Tile Regions, click on Verify and click on Set Z &#38; move to next.</li>
		<li>Under the Focus Strategy tab, check that the focus strategy is set to Use Z Values/Focus Surface defined in the Tiles Setup.</li>
		<li>Click on Start Experiment to start the image acquisition.</li>
	</ol>
	</li>
	<li>Image processing
	<ol>
		<li>In the Processing tab, select Single or Batch mode. Select Airyscan processing. Tick 3D processing and select Standard Autofilter. Select the&#160;raw image acquired and start processing (Supplemental Figure S2A).</li>
		<li>Once the Airyscan Processing is completed, select Stitching in the processing methods. Select New Output, Fuse Tiles, Correct Shading, and All by reference, with the reference as DAPI. Select the Airyscan processed image and start processing (Supplemental Figure S2B).</li>
		<li>Save the raw and double-processed images. The latter is saved automatically in Batch mode.</li>
	</ol>
	</li>
</ol>
6. Image analysis
NOTE: The analysis pipeline used here was customized to the purposes of this study. The details of the pipeline operations are shown in Supplemental Figure S3.
<ol>
	<li>Import the czi image file into the software. Open the analysis panel; in the dropdown select New Pipeline&#8230;</li>
	<li>To create a cell detection pipeline, import the Cellpose Python Segmenter into the pipeline12. Set Model_Name to cyto2, Input_channel to 2, Second_channel to 3, Diameter_in_&#181;m to 10, Flow_threshold to 0.4, and Cellprob_threshold to 0. Add an Import Document Objects (renamed to Import&#160;Manual Labels) before and an Object Feature Filter (renamed to Size Filter) operator after the pipeline by clicking on + Add Operation. See Supplemental Figure S3A for the values of the pipeline settings.
	<ol>
		<li>Move viewer to the first plane of interest</li>
		<li>Click on the Draw Objects Tool icon</li>
		<li>Click on the Polygon mode icon</li>
		<li>Draw area to analyzed</li>
		<li>Move viewer to the last plane of interest</li>
		<li>Draw area to be analyzed</li>
		<li>Click on the green tick Apply Changes</li>
		<li>Click on the Show Objects Table icon</li>
		<li>Right-click on the created object and select Rename Object to rename the object to an appropriate name. Right-click on the created object and click on Add Tag. Add a tag called Analysis Region.</li>
	</ol>
	</li>
	<li>Run this pipeline by clicking on the forward blue arrow at the top of the Analysis panel.</li>
	<li>To create a primary cilia and centriole detection pipeline, open the analysis panel, in the drop down select New Pipeline&#8230;. Add the following operators to the pipeline: Import Document Objects (renamed to Import Manual Labels), two Intensity Threshold Segmenter&#160;operators (renamed to Cilia Threshold and Centriole Threshold), Splitting (renamed to Centriole Splitting), Object Feature Filter (renamed to Centriole Splitting size filter) and Compartment (renamed to Compartment - Cell). See Supplemental Figure S3B for the values of the pipeline settings.</li>
	<li>Run this pipeline by clicking on the forward blue arrow at the top of the Analysis panel.</li>
	<li>Open the Objects window. Click on Im/Export&#8230;, select Excel Export&#8230;. Select the following features to save: # Children, Name, Parent Names, 3D oriented bounds Short side, Middle side, Long side, Angle XY, Angle XZ, Angle YZ, Bounding Box X1, X2, Y1, Y2, Z1, Z2, SizeX, SizeY, SizeZ, Center of Geometry X, Y, Z, Plane First, Last, Count, Sphericity (Mesh), Volume (Mesh), Surface Area (Mesh).</li>
</ol>

<ol>
	<li>Wheatley, D. N., Wang, A. M., Strugnell, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Expression+of+primary+cilia+in+mammalian+cells.">Expression of primary cilia in mammalian cells.</a> Cell Biol Int. 20 (1), 73-81 (1996).</li><li>Donnelly, E., Ascenzi, M. G., Farnum, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primary+cilia+are+highly+oriented+with+respect+to+collagen+direction+and+long+axis+of+extensor+tendon.">Primary cilia are highly oriented with respect to collagen direction and long axis of extensor tendon.</a> J Orthop Res. 28 (1), 77-82 (2010).</li><li>McGlashan, S. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+loading+modulates+chondrocyte+primary+cilia+incidence+and+length.">Mechanical loading modulates chondrocyte primary cilia incidence and length.</a> Cell Biol Int. 34 (5), 441-446 (2010).</li><li>Farnum, C. E., Wilsman, N. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Orientation+of+primary+cilia+of+articular+chondrocytes+in+three-dimensional+space.">Orientation of primary cilia of articular chondrocytes in three-dimensional space.</a> Anat Rec. 294 (3), 533-549 (2011).</li><li>Coveney, C. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+ciliary+protein+IFT88+controls+post-natal+cartilage+thickness+and+influences+development+of+osteoarthritis.">The ciliary protein IFT88 controls post-natal cartilage thickness and influences development of osteoarthritis.</a> Arth Rheumatol. 74 (1), 49-59 (2021).</li><li>Coveney, C. R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Ciliary+IFT88+protects+coordinated+adolescent+growth+plate+ossification+from+disruptive+physiological+mechanical+forces.">Ciliary IFT88 protects coordinated adolescent growth plate ossification from disruptive physiological mechanical forces.</a> J Bone Miner Res. 37 (6), 1081-1086 (2022).</li><li>Andrea, C. E. D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Primary+cilia+organization+reflects+polarity+in+the+growth+plate+and+implies+loss+of+polarity+and+mosaicism+in+osteochondroma.">Primary cilia organization reflects polarity in the growth plate and implies loss of polarity and mosaicism in osteochondroma.</a> Lab Invest. 90 (7), 1091-1101 (2010).</li><li>Ascenzi, M. -. G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effect+of+localization,+length+and+orientation+of+chondrocytic+primary+cilium+on+murine+growth+plate+organization.">Effect of localization, length and orientation of chondrocytic primary cilium on murine growth plate organization.</a> J. Theor Biol. 285 (1), 147-155 (2011).</li><li>Bangs, F. K., Schrode, N., Hadjantonakis, A. K., Anderson, K. V. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lineage+specificity+of+primary+cilia+in+the+mouse+embryo.">Lineage specificity of primary cilia in the mouse embryo.</a> Nat. Cell Biol. 17 (2), 113-122 (2015).</li><li>Larkins, C. E., Aviles, G. D. G., East, M. P., Kahn, R. A., Caspary, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Arl13b+regulates+ciliogenesis+and+the+dynamic+localization+of+Shh+signaling+proteins.">Arl13b regulates ciliogenesis and the dynamic localization of Shh signaling proteins.</a> Mol Biol Cell. 22 (23), 4694-4703 (2011).</li><li>Higginbotham, H., Bielas, S., Tanaka, T., Gleeson, J. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transgenic+mouse+line+with+green-fluorescent+protein-labeled+Centrin+2+allows+visualization+of+the+centrosome+in+living+cells.">Transgenic mouse line with green-fluorescent protein-labeled Centrin 2 allows visualization of the centrosome in living cells.</a> Transgenic Res. 13 (2), 155-164 (2004).</li><li>arivis. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Application Note #47 - Applying Cellpose models in arivis Vision4D. , 1-20 (2021).</li><li>Huff, J., Bergter, A., Luebbers, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Application+Note:+Multiplex+mode+for+the+LSM+9+series+with+Airyscan+2:+fast+and+gentle+confocal+super-resolution+in+large+volumes.">Application Note: Multiplex mode for the LSM 9 series with Airyscan 2: fast and gentle confocal super-resolution in large volumes.</a> Nat Methods. , (2019).</li><li>Ning, K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cilia-associated+wound+repair+mediated+by+IFT88+in+retinal+pigment+epithelium.">Cilia-associated wound repair mediated by IFT88 in retinal pigment epithelium.</a> Sci Rep. 13 (1), 8205-8219 (2023).</li><li>Palla, A. 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The authors have no conflicts of interest to declare.

In this protocol, it is essential to proceed with the tissue processing and embedding steps immediately after collection. The blocks and slides of tissue, however, can be kept for at least 3 months at -20 &#176;C. Several steps in this protocol can be modified to suit different applications, in particular, section thickness, magnification, and z-stack parameters. This will impact the depth of imaging and the resolution obtained. Changes in numerical aperture and resolution mode could be used to obtain an even higher resolution and the z-stacks interval size decreased. However, this would increase imaging time or reduce the size of the field of view that can be imaged. The imaging parameters described are the minimum requirements identified for 3D analysis of primary cilia orientation.
The choice of mounting medium is important to obtain high resolution. The refractive index of the mounting medium must be similar to that of the immersion medium to minimize image degradation in the z-stack. The refractive index of the mountant and the immersion oil used here have refractive indexes of 1.52 and 1.518, respectively. The Airyscan function on the confocal microscope used here offers super-resolution images. Other super-resolution microscopes could be used; however, we found that the confocal mode only was not sufficient to obtain the resolution required for the size of the areas imaged. The maximum resolution of the Multiplex SR-8Y Airyscan mode is 120/160 nm in x/y and 450 nm in z, and a maximum number of frames/s of 47.5. We found the Multiplex SR-4Y Airyscan mode, which presents a maximum resolution of 140 nm in x/y and 450 nm in z but images at half the speed with a maximum number of frames/s of 25, not to present a significant difference for imaging primary cilia in 3D. Similarly, the Airyscan super-resolution (SR) mode, which offers the highest resolution on this microscope -- 120 nm in x/y and 350 nm in z -- would take longer to image, as its maximum number of frames/s is 4.713.
Background GFP signal can be observed in other tissues such as the muscle. To minimize this, the range of the lasers can be narrowed to reduce overlap with other wavelengths. This line has previously been used to image primary cilia in the embryo, primary cell cultures, muscle fibers, and the retinal pigment epithelium9,14,15. We have successfully imaged primary cilia in kidney, cartilage, and bone tissue. Some optimization of imaging settings will likely be required for different tissues; however, this cryosection protocol should ensure preservation of the signal.
Two-cell thick sections (40-60 &#181;m) sections are sufficient to obtain 3D orientation information in the growth plate, as this allows imaging of whole cells and their primary cilia. For other tissues where cells are larger than chondrocytes, such as adipocytes, thicker sections may be required to collect cell population data. Depending on the depth resolution of the microscope used, it may not be possible to image deep into thicker sections. Tissue clearing techniques would likely lead to loss of the endogenous mCherry and GFP signal due to the time it requires. This was explored, as clearing methods were used for other imaging in the limb but it was not found necessary in this context. Our experience with endogenous CD31-RFP suggested clearing techniques would need careful, bespoke to sample, optimization.
This ARL13B-CENTRIN-2 line can be crossed with other mouse lines to further study the organization of primary cilia in tissues. For example, we have crossed it with an aggrecan-Cre IFT88fl/fl to conditionally knock out IFT88, a primary cilia protein whose deletion prevents ciliation, in aggrecan-expressing cells (postnatal chondrocytes)5,6. These ARL13B and CENTRIN-2 fluorescent signals can then be used to measure the efficiency of genetic perturbations at the level of ciliary structure in tissues and any associated changes in ciliary organization. Ongoing work is currently doing exactly this. Our first aim will be to quantify, with this more robust methodology, the effect of IFT88 deletion on cilia prevalence. Previously, we have gauged this by using immunofluorescence and estimated this to be a 20% reduction6. This method can also be combined with immunohistochemistry to image primary cilia in combination with other proteins, for example, a cell membrane protein.
The transgene on ARL13B may have an effect on ARL13B expression and primary cilia length. Previous studies have measured different primary cilia lengths with the ARL13B-CENTRIN-2 line compared to non-transgene ARL13B in certain regions of the mouse brain, with no difference in others16. Another study observed differences in primary cilia length in a GFP-tagged ARL13B mouse line compared to untagged ARL13B in mouse embryonic fibroblasts10. Ongoing work is verifying any changes in primary cilia length in the growth plate and other tissues by comparing the results using the ARL13B-CENTRIN-2 mouse line with antibody staining against non-transgenic ARL13B.
The image analysis pipeline in this protocol was designed to subsequently measure the ciliation percentage, length, and 3D orientation of primary cilia, as well as centriole number and position in the growth plate. The image analysis software allows for custom pipelines to be made by changing the steps involved. The output of this pipeline is a spreadsheet with the measures of the chosen features for the cells, cilia, and centrioles detected. Postanalysis can then be performed in R or other software according to the purposes of the study. As with the imaging settings, the pipeline steps and specific settings can be changed to work on different images and according to the aims of the study. Overall, this method allows high-throughput analysis of primary cilia than has been published previously in the growth plate. The number of cells and associated cilia that can be studied allows for larger and more reliable measurements.

The primary cilium is an essential organelle in the development and homeostasis of many tissues and organs, including the skeleton. During postnatal growth, it regulates cartilage mineralization during endochondral ossification1. In humans, genetic mutations that affect cilia-related proteins cause a range of diseases and developmental disorders, known as ciliopathies. An understanding of the organization of primary cilia inside tissues is important in understanding how they regulate cellular behavior at the organ level. Incidence, length, and orientation of primary cilia have been shown to vary within and between tissues, suggestive of a role for tissue organization to their function2,3,4. The growth plate, the layer of cartilage at the end of long bones where most postnatal bone growth occurs from early life and through adolescence, is a highly organized tissue with chondrocyte cells arranged in columns. In the growth plate, chondrocytes go through stages of differentiation, from the resting to proliferating to hypertrophic.
This cellular organization is lost in some ciliopathies, which suggests primary cilia may play a role in coordinating this cellular organization and/or, at the very least, be associated with cellular polarization. In addition, primary cilia orientation appears to play a role in their mechanosensory function, with differences observed between load-bearing and non-load-bearing regions in articular cartilage4. Our own recent studies have implied differential roles within different regions of articular cartilage and the growth plate5,6. Previous studies that have imaged primary cilia in the growth plate have been limited to 2D, which loses information related to cilia organization in tissue or are limited to a small number of cells in 3D, which reduces the scale and power of the analysis7,8.
Among the challenges to imaging the primary cilium are its micrometer-scale size and the non-specificity of immunohistochemistry antibodies. Using a native fluorescent signal ensures strong localization and specificity. Native fluorescent signals can, however, be challenging to image in mineralized tissues, as the long decalcification process to embed tissues in paraffin can result in loss of signal. Cryosectioning samples frozen in the native state prevents this loss by speeding up the embedding and sectioning process.
To image primary cilia in situ, a double transgenic mouse line was generated by Bangs et al., which expresses ADP-ribosylation factor-like protein (ARL13B) fused to mCherry and CENTRIN-2 fused to GFP9. ARL13B is a small GTPase localized to the primary cilium membrane, and CENTRIN-2 is a centriolar protein localized to the two centrosomes at the base of the cilium10,11. This mouse line, therefore, allows fluorescently labeled primary cilia to be imaged without the need for immunohistochemistry.
The protocol described here makes use of this transgenic line to image primary cilia in 3D in murine postnatal growth plates with confocal microscopy, only requiring additional 4&#39;,6-diamidino-2-phenylindole (DAPI) staining. Hundreds of primary cilia in hundreds of cells can be imaged and analyzed collectively in a tissue context. An image analysis pipeline subsequently measures primary cilia incidence, length, and orientation in the different regions of the growth plate. The pipeline is also able to quantify these features for centrioles. An outline of the steps in the protocol is presented in Figure 1.

<table><tbody><tr><td>22 x 50 mm cover slips</td><td>fisherscientific&amp;nbsp;</td><td>12373128</td><td></td></tr><tr><td>arivis Vision4D software</td><td>Zeiss</td><td>N/A</td><td>Image analysis software&amp;nbsp;</td></tr><tr><td>Confocal laser scanning microscope&amp;nbsp;</td><td>Zeiss</td><td>Zeiss 980 Airyscan 2</td><td></td></tr><tr><td>CoverGrip Coverslip Sealant&amp;nbsp;</td><td>Biotium&amp;nbsp;</td><td>23005</td><td></td></tr><tr><td>Cryofilm type 3C(16UF) (sheet type) 2.0 cm width</td><td>Section-lab</td><td>C-FUF303</td><td>To be purchased from Section-lab directly</td></tr><tr><td>Cryostat</td><td>Leica</td><td>CM1900 UV</td><td></td></tr><tr><td>DAPI</td><td>invitrogen</td><td>D1306</td><td></td></tr><tr><td>Disposable Base Molds&amp;nbsp;</td><td>epredia</td><td>58952</td><td></td></tr><tr><td>Dulbecco&#039;s Phosphate Buffered Saline</td><td>gibco</td><td>14190</td><td></td></tr><tr><td>Formalin</td><td>Sigma</td><td>20260630</td><td></td></tr><tr><td>Imaris software&amp;nbsp;</td><td>Oxford Instruments&amp;nbsp;</td><td>N/A</td><td>Image analysis software for 3D reconstruction&amp;nbsp;</td></tr><tr><td>Immersion Oil 518F</td><td>Zeiss</td><td>ISO 8036&amp;nbsp;</td><td></td></tr><tr><td>Microtome blades</td><td>Feather</td><td>02.075.00.006</td><td></td></tr><tr><td>SlowFade Glass Antifade Mountant&amp;nbsp;</td><td>invitrogen</td><td>S36917</td><td></td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S9378</td><td></td></tr><tr><td>Super Cryoembedding Medium (SCEM)</td><td>Section-lab</td><td>C-EM001</td><td>To be purchased from Section-lab directly</td></tr><tr><td>Superfrost Plus Microscope slides</td><td>avantor</td><td>631-0108</td><td></td></tr><tr><td>Zen software</td><td>Zeiss</td><td>N/A</td><td>Freeware for Confocal laser scanning microscopy</td></tr></tbody></table>

volumetric imaging and analysis of primary cilia in musculoskeletal tissue using the arl13b-centrin-2 mouse model

The primary cilium is a non-motile, solitary organelle assembled by most cell types. It is essential to the development and homeostasis of the skeleton and in coordinating cell responses to biochemical and mechanical cues from their surrounding environment. Due to its micrometer-scale size, the primary cilium is challenging to image in situ to characterize in tissues, in a high throughput manner and without optical biases. An understanding of its organization in its 3D environment in vivo will be key to understanding ciliary roles in physiology and disease. The ARL13B-CENTRIN-2 mouse line presents an mCherry tag on the ciliary axoneme protein ARL13B and a green fluorescent protein (GFP) tag on the centriole protein CENTRIN-2, localized to the base of the cilium. This mouse line allows for imaging of the primary cilium without the need for staining, which reduces the number of steps in image collection and circumnavigates issues with signal-to-noise. This publication describes a step-by-step protocol to image primary cilia in musculoskeletal tissues whilst preserving the fluorescent signal. It also describes a universally applicable image analysis pipeline, enabling unbiased quantification of cilia features, such as length and 3D orientation.

The growth plate of 6-week-old mouse was imaged using the described protocol and primary cilia organization mapped in the tissue. An image of all three regions of the growth plate -- resting, proliferative, and hypertrophic -- was captured and run through the analysis pipeline (Figure 2A). Individual cells can be identified with the DAPI staining, with their ciliary axoneme in red and their centrioles in green. The resolution using this method is high enough to discriminate individual centrioles (Figure 2B). At this resolution, the cilia can be visualized in their 3D environment, pointing in different directions (Video 1).
Lower resolution images, with higher magnification objectives, can also be captured to visualize primary cilia across a larger region of tissue; however, these are not suitable for analysis of cilia orientation in 3D. For instance, the primary cilia in articular cartilage were imaged with the 20x objective, demonstrating that the method also works on other tissues (Figure 2C). At this resolution, the ciliary axonemes in red can be identified, but the centrioles are more difficult to discriminate. Subchondral bone tends to present a lot of GFP background signal, making the identification of centriole difficult in this tissue.
The pipeline can detect individual cells, primary cilia, and centrioles, and it works for highly organized young growth plates and older growth plates where the columns are less well defined, as in 10-week-old mice (Figure 3). Cells and cilia are detected in 3D, as can be viewed across the z-stack.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67693/67693fig01.jpg" /> 
Figure 1: Experimental workflow. Flowchart of the different steps in the protocol from tissue collection to image analysis. <a href="https://www.jove.com/files/ftp_upload/67693/67693fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67693/67693fig02.jpg" /> 
Figure 2: Primary cilia in murine tissues of the ARL13B-CENTRIN-2 mouse. (A) A 6-week-old growth plate, with DAPI in white, CENTRIN-2 in green, and ARL13B in red, imaged with the 40x objective. (B) Zoom in on an individual primary cilium and its two centrioles. (C) Articular cartilage and subchondral bone of a 4-week-old mouse, imaged with the 20x objective. (D) Kidney of a 6-week-old mouse imaged using the same pipeline as for musculoskeletal tissue. (E) Magnification of the white box in D showing individual primary cilia and their centrioles in the kidney. Scale bars = 20 &#181;m (A,D), 4 &#181;m (B), 60 &#181;m (C), 6 &#181;m (E). <a href="https://www.jove.com/files/ftp_upload/67693/67693fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/67693/67693fig03.jpg" /> 
Figure 3: Cell and cilia detection pipeline. Example application of the image analysis pipeline on a 10-week-old growth plate. (A) Image opened in the software. (B) Result of the cell detection pipeline. (C) Result of the cilia and centriole detection pipeline. Cilia are in pink and centrioles in green. The side view exemplifies that the resolution is high enough to discriminate cilia oriented in different directions in 3D. <a href="https://www.jove.com/files/ftp_upload/67693/67693fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Video 1: Three-dimensional reconstruction of a 6-week-old growth plate from the ARL13B-CENTRIN-2 mouse. The three-dimensional reconstruction was conducted in the Imaris software and recorded as an animation video. <a href="https://www.jove.com/files/ftp_upload/67693/Video-1.mp4" target="_blank">Please click here to download this Video.</a>
Supplemental Figure S1: Image acquisition settings on the microscope system. (A) Smart Setup window to add the channels and choose the Airyscan mode. (B) Tile viewer to select and preview the tiles to image. (C) Airyscan Detector Adjustment window, where the alignment of the Airyscan detectors is confirmed once all the tiles turn green. (D) Laser settings for the three channels. (E) Acquisition Mode and Z-Stack settings. <a href="https://www.jove.com/files/ftp_upload/67693/Supp-file-1.zip" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Image processing settings on the microscope system. (A) Airyscan Processing settings. (B) Stitching settings. <a href="https://www.jove.com/files/ftp_upload/67693/Supp-file-2.zip" target="_blank">Please click here to download this File.</a>
Supplemental Figure S3: Custom image analysis pipelines. (A) Cell detection pipeline using the GFP and DAPI channels. (B) Primary cilia and centriole detection pipeline using the mCherry and GFP channels, respectively. <a href="https://www.jove.com/files/ftp_upload/67693/Supp-file-3-.zip" target="_blank">Please click here to download this File.</a>

This method describes the exploitation of the endogenous signal in an ARL13B-CENTRIN-2 transgenic mouse model to image primary cilia in situ, in three dimensions across large areas of mineralized tissue, from tissue collection to analysis of cilia organization. This protocol may also be applied to other types of tissues.

Volumetric Imaging and Analysis of Primary Cilia in Musculoskeletal Tissue using the ARL13B-CENTRIN-2 Mouse Model

The primary cilium is a non-motile, solitary organelle assembled by most cell types. It is essential to the development and homeostasis of the skeleton ...

volumetric-imaging-analysis-primary-cilia-musculoskeletal-tissue

Research

JoVE Journal

Biology

319 Views.  University of Oxford. Deze methode beschrijft de exploitatie van het endogene signaal in een ARL13B-CENTRIN-2 transgeen muismodel om primaire cilia in situ in beeld te brengen, in drie dimensies over grote delen van gemineraliseerd weefsel, van weefselverzameling tot analyse van de organisatie van cilia's. Dit protocol kan ook worden toegepast op andere soorten weefsels. 

Video: Volumetrische beeldvorming en analyse van primaire trilharen in musculoskeletaal weefsel met behulp van het ARL13B-CENTRIN-2-muismodel

In situ Imaging of the Mouse Thymus Using 2-Photon Microscopy

Two-photon Microscopy (TPM) enables us to image deep into the thymus and document the events that are important for thymocyte development. To follow the migration of individuals in a crowd of thymocytes , we generate neonatal chimeras where less than one percent of the thymocytes are derived from a donor that is transgenic for a ubiquitously express fluorescent protein. To generate these partial hematopoetic chimeras, neonatal recipients are injected with bone marrow between 3-7 days of age. After 4-6 weeks, the mouse is sacrificed and the thymus is carefully dissected and bissected preserving the architecture of the tissue that will be imaged. The thymus is glued onto a coverslip in preparation for ex vivo imaging by TPM. During imaging the thymus is kept in DMEM without phenol red that is perfused with 95% oxygen and 5% carbon dioxide and warmed to 37&deg;C. Using this approach, we can study the events required for the generation of a diverse T cell repertoire.

We present step-by-step instructions for the generation of neonatal chimeras as well as the dissection and preparation of the thymus for ex vivo imaging by 2-Photon Microscopy.

Two-photon Microscopy (TPM) enables us to image deep into the thymus and document the events that are important for thymocyte development.  To follow the ...

Actin Co-Sedimentation Assay; for the Analysis of Protein Binding to F-Actin

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates tension and helps maintains cellular shape.  Although the actin cytoskeleton is a rigid structure, it is a dynamic structure that is constantly remodeling.  A number of proteins can bind to the actin cytoskeleton.  The binding of a particular protein to F-actin is often desired to support cell biological observations or to further understand dynamic processes due to remodeling of the actin cytoskeleton. The actin co-sedimentation assay is an in vitro assay routinely used to analyze the binding of specific proteins or protein domains with F-actin.  The basic principles of the assay involve an incubation of the protein of interest (full length or domain of) with F-actin, ultracentrifugation step to pellet F-actin and analysis of the protein co-sedimenting with F-actin.  Actin co-sedimentation assays can be designed accordingly to measure actin binding affinities and in competition assays.

Proteins bind to filamentous actin (F-actin) through distinct actin binding modules. In this video we demonstrate the procedure of actin co-sedimentation, which is an in vitro assay routinely used to analyze proteins or specific domains that bind F-actin.

The actin cytoskeleton within the cell is a network of actin filaments that allows the movement of cells and cellular processes, and that generates ...

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Analysis of Gene Function and Visualization of Cilia-Generated Fluid Flow in Kupffer's Vesicle

Internal organs such as the heart, brain, and gut develop left-right (LR) asymmetries that are critical for their normal functions1. Motile cilia are involved in establishing LR asymmetry in vertebrate embryos, including mouse, frog, and zebrafish2-6. These 'LR cilia' generate asymmetric fluid flow that is necessary to trigger a conserved asymmetric Nodal (TGF-&beta; superfamily) signaling cascade in the left lateral plate mesoderm, which is thought to provide LR patterning information for developing organs7. Thus, to understand mechanisms underlying LR patterning, it is essential to identify genes that regulate the organization of LR ciliated cells, the motility and length of LR cilia and their ability to generate robust asymmetric flow.
In the zebrafish embryo, LR cilia are located in Kupffer's vesicle (KV)2,4,5. KV is comprised of a single layer of monociliated epithelial cells that enclose a fluid-filled lumen. Fate mapping has shown that KV is derived from a group of ~20-30 cells known as dorsal forerunner cells (DFCs) that migrate at the dorsal blastoderm margin during epiboly stages8,9. During early somite stages, DFCs cluster and differentiate into ciliated epithelial cells to form KV in the tailbud of the embryo10,11. The ability to identify and track DFCs&mdash;in combination with optical transparency and rapid development of the zebrafish embryo&mdash;make zebrafish KV an excellent model system to study LR ciliated cells.
Interestingly, progenitors of the DFC/KV cell lineage retain cytoplasmic bridges between the yolk cell up to 4 hr post-fertilization (hpf), whereas cytoplasmic bridges between the yolk cell and other embryonic cells close after 2 hpf8. Taking advantage of these cytoplasmic bridges, we developed a stage-specific injection strategy to deliver morpholino oligonucleotides (MO) exclusively to DFCs and knockdown the function of a targeted gene in these cells12. This technique creates chimeric embryos in which gene function is knocked down in the DFC/KV lineage developing in the context of a wild-type embryo. To analyze asymmetric fluid flow in KV, we inject fluorescent microbeads into the KV lumen and record bead movement using videomicroscopy2. Fluid flow is easily visualized and can be quantified by tracking bead displacement over time.
Here, using the stage-specific DFC-targeted gene knockdown technique and injection of fluorescent microbeads into KV to visualize flow, we present a protocol that provides an effective approach to characterize the role of a particular gene during KV development and function.

Cilia-generated fluid flow in Kupffer&#x2019;s Vesicle (KV) controls left-right patterning of the zebrafish embryo. Here, we describe a technique to modulate gene function specifically in KV cells. In addition, we show how to deliver fluorescent beads into KV to visualize fluid flow.

Internal organs such as the  heart, brain, and gut develop left-right (LR) asymmetries that are critical for  their normal functions1. Motile cilia are ...

Isolation of Primary Myofibroblasts from Mouse and Human Colon Tissue

The myofibroblast is a stromal cell of the gastrointestinal (GI) tract that has been gaining considerable attention for its critical role in many GI functions. While several myofibroblast cell lines are commercially available to study these cells in vitro, research results from a cell line exposed to experimental cell culture conditions have inherent limitations due to the overly reductionist nature of the work. Use of primary myofibroblasts offers a great advantage in terms of confirming experimental findings identified in a cell line. Isolation of primary myofibroblasts from an animal model allows for the study of myofibroblasts under conditions that more closely mimic the disease state being studied. Isolation of primary myofibroblasts from human colon tissue provides arguably the most relevant experimental data, since the cells come directly from patients with the underlying disease. We describe a well-established technique that can be utilized to isolate primary myofibroblasts from both mouse and human colon tissue. These isolated cells have been characterized to be alpha-smooth muscle actin and vimentin-positive, and desmin-negative, consistent with subepithelial intestinal myofibroblasts. Primary myofibroblast cells can be grown in cell culture and used for experimental purposes over a limited number of passages.

The myofibroblast is an influential stromal cell of the gastrointestinal tract that regulates important physiologic processes in both normal and disease states. We describe a technique that allows for the isolation of primary myofibroblasts from both mouse and human colon tissue, which can be utilized for in vitro experimentation.

The myofibroblast is  a stromal cell of the gastrointestinal (GI) tract that has been gaining  considerable attention for its critical role in many GI ...

High-resolution Time-lapse Imaging and Automated Analysis of Microtubule Dynamics in Living Human Umbilical Vein Endothelial Cells

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes within individual endothelial cells (ECs). These processes are critical for homeostatic maintenance such as wound healing, and are also crucial in promoting tumor growth and metastasis. EC morphology is defined by the organization of the cytoskeleton, a tightly regulated system of actin and microtubule (MT) dynamics that is known to control EC branching, polarity and directional migration, essential components of angiogenesis. To study MT dynamics, we used high-resolution fluorescence microscopy coupled with computational image analysis of fluorescently-labeled MT plus-ends to investigate MT growth dynamics and the regulation of EC branching morphology and directional migration. Time-lapse imaging of living Human Umbilical Vein Endothelial Cells (HUVECs) was performed following transfection with fluorescently-labeled MT End Binding protein 3 (EB3) and Mitotic Centromere Associated Kinesin (MCAK)-specific cDNA constructs to evaluate effects on MT dynamics. PlusTipTracker software was used to track EB3-labeled MT plus ends in order to measure MT growth speeds and MT growth lifetimes in time-lapse images. This methodology allows for the study of MT dynamics and the identification of how localized regulation of MT dynamics within sub-cellular regions contributes to the angiogenic processes of EC branching and migration.

Protocols for Human Umbilical Vein Endothelial Cell (HUVEC) culture, transient transfection of fluorescently-labeled markers of microtubule growth, live-cell imaging and automated analysis of interphase microtubule growth dynamics are detailed.

The physiological process by which new vasculature forms from existing vasculature requires specific signaling events that trigger morphological changes ...

Dissection of the Auditory Bulla in Postnatal Mice: Isolation of the Middle Ear Bones and Histological Analysis

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called the auditory bulla houses the ossicles, whereas the auditory capsule encloses the inner ear, namely the cochlea and semicircular canals. Murine ossicles are essential for hearing and thus of great interest to researchers in the field of otolaryngology, but their metabolism, development, and evolution are highly relevant to other fields. Altered bone metabolism can affect hearing function in adult mice, and various gene-deficient mice show changes in morphogenesis of auditory ossicles in utero. Although murine auditory ossicles are tiny, their manipulation is feasible if one understands their anatomical orientation and 3D structure. Here, we describe how to dissect the auditory bulla and capsule of postnatal mice and then isolate individual ossicles by removing part of the bulla. We also discuss how to embed the bulla and capsule in different orientations to generate paraffin or frozen sections suitable for preparation of longitudinal, horizontal, or frontal sections of the malleus. Finally, we enumerate anatomical differences between mouse and human auditory ossicles. These methods would be useful in analyzing pathological, developmental and evolutionary aspects of auditory ossicles and the middle ear in mice.

We present a protocol to isolate the auditory bulla, capsule, and ossicles from postnatal mice for whole mount and histological analysis.

In most mammals, auditory ossicles in the middle ear, including the malleus, incus and stapes, are the smallest bones. In mice, a bony structure called ...

Analysis of Beta-cell Function Using Single-cell Resolution Calcium Imaging in Zebrafish Islets

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to hyperglycemia and severe, life-threatening consequences. Understanding how the beta-cells operate under physiological conditions and what genetic and environmental factors might cause their dysfunction could lead to better treatment options for diabetic patients. The ability to measure calcium levels in beta-cells serves as an important indicator of beta-cell function, as the influx of calcium ions triggers insulin release. Here we describe a protocol for monitoring the glucose-stimulated calcium influx in zebrafish beta-cells by using GCaMP6s, a genetically encoded sensor of calcium. The method allows monitoring the intracellular calcium dynamics with single-cell resolution in ex vivo mounted islets. The glucose-responsiveness of beta-cells within the same islet can be captured simultaneously under different glucose concentrations, which suggests the presence of functional heterogeneity among zebrafish beta-cells. Furthermore, the technique provides high temporal and spatial resolution, which reveals the oscillatory nature of the calcium influx upon glucose stimulation. Our approach opens the doors to use the zebrafish as a model to investigate the contribution of genetic and environmental factors to beta-cell function and dysfunction.

Beta-cell functionality is important for blood-glucose homeostasis, which is evaluated at single-cell resolution using a genetically encoded reporter for calcium influx.

Pancreatic beta-cells respond to increasing blood glucose concentrations by secreting the hormone insulin. The dysfunction of beta-cells leads to ...

Adipo-Clear: A Tissue Clearing Method for Three-Dimensional Imaging of Adipose Tissue

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte precursors, immune cells, fibroblasts, blood vessels, and nerve projections. Although the molecular control of cell type specification and how these cells interact have been increasingly delineated, a more comprehensive understanding of these adipose-resident cells can be achieved by visualizing their distribution and architecture throughout the whole tissue. Existing immunohistochemistry and immunofluorescence approaches to analyze adipose histology rely on thin paraffin-embedded sections. However, thin sections capture only a small portion of tissue; as a result, the conclusions can be biased by what portion of tissue is analyzed. We have therefore developed an adipose tissue clearing technique, Adipo-Clear, to permit comprehensive three-dimensional visualization of molecular and cellular patterns in whole adipose tissues. Adipo-Clear was adapted from iDISCO/iDISCO+, with specific modifications made to completely remove the lipid stored in the tissue while preserving native tissue morphology. In combination with light-sheet fluorescence microscopy, we demonstrate here the use of the Adipo-Clear method to obtain high-resolution volumetric images&#160;of an entire adipose tissue.

Due to the high lipid content, adipose tissue has been challenging to visualize using traditional histological methods. Adipo-Clear is a tissue clearing technique that allows robust labeling and high-resolution volumetric fluorescent imaging of adipose tissue. Here, we describe the methods for sample preparation, pretreatment, staining, clearing, and mounting for imaging.

Adipose tissue plays a central role in energy homeostasis and thermoregulation. It is composed of different types of adipocytes, as well as adipocyte ...

Measurement of Energy Metabolism in Explanted Retinal Tissue Using Extracellular Flux Analysis

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while maintaining transparency of the visual axis. Perturbations to this delicate balance cause blinding illnesses, such as diabetic retinopathy. Therefore, the understanding of energy metabolism changes in the retina during disease is imperative to the development of rational therapies for various causes of vison loss. The recent advent of commercially-available extracellular flux analyzers has made the study of retinal energy metabolism more accessible. This protocol describes the use of such an analyzer to measure contributions to retinal energy supply through its two principle arms - oxidative phosphorylation and glycolysis - by quantifying changes in oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) as proxies for these pathways. This technique is readily performed in explanted retinal tissue, facilitating assessment of responses to multiple pharmacologic agents in a single experiment. Metabolic signatures in retinas from animals lacking rod photoreceptor signaling are compared to wild-type controls using this method. A major limitation in this technique is the lack of ability to discriminate between light-adapted and dark-adapted energy utilization, an important physiologic consideration in retinal tissue.

This technique describes real time recording of oxygen consumption and extracellular acidification rates in explanted mouse retinal tissues using an extracellular flux analyzer.

High acuity vision is a heavily energy-consuming process, and the retina has developed several unique adaptations to precisely meet such demands while ...

Volumetric Imaging and Analysis of Primary Cilia in Musculoskeletal Tissue using the ARL13B-CENTRIN-2 Mouse Model

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