This work was supported by the School of Medicine and The Institute for Obesity Research of Tecnologico de Monterrey. Figure 3A was created with Scientific Image and Illustration software.

All animal experimentation procedures were evaluated and approved by the Animal Use and Care Committee (CICUAL) of Tecnologico de Monterrey (Protocol 2019-007). Male Zucker (+/+ and fa/fa) rats aged 12 to 13 weeks were used for this study. Animals were kept in standard husbandry conditions (12 h/12 h light/dark cycle, 40-60% humidity) and had access to food (standard rat chow) and water ad libitum.
1. Solution composition
<ol>
	<li>Prepare fresh Relax solution by mixing the components shown in Table 1. Adjust pH to 7.3 using sodium hydroxide (NaOH).</li>
	<li>Calculate the free calcium concentration in the Relax solution using a simulation program for determining the free metal concentration.</li>
	<li>Prepare a 5 mM tetramethylrhodamine methyl ester (TMRE) stock in dimethyl sulfoxide (DMSO) and a subsequent dilution of 0.1 mM in DMSO.</li>
</ol>
2. Dissection of the gastrocnemius muscle fiber bundles
<ol>
	<li>Place the animal inside the induction chamber and close the lid. Turn the oxygen source on, set the gas flow rate at 0.5 L/min, and set the vaporizer at 4% sevoflurane to induce anesthesia.</li>
	<li>Once the rat is asleep, place it outside the chamber in a supine position while maintaining anesthesia. Pinch the toe or tail to verify the lack of reflexes.</li>
	<li>With scissors, cut through the abdomen&#39;s skin and muscles. Then, cut the thoracic cage to gain access to the heart.</li>
	<li>To perform a cardiectomy, grasp the heart from the topmost veins and arteries with forceps and cut the blood vessels with scissors. Remove the heart.</li>
	<li>Immediately after euthanasia, disinfect the hindlimb with ethanol and shave it using a shaving machine.</li>
	<li>Hold the hindfoot and make an incision with scissors through the skin at the level of the Achilles tendon.</li>
	<li>Cut the hindlimb with scissors at the level of the proximal tibia. Transfer it to a 60 mm Petri dish containing 3 mL of ice-cold Relax solution in a dorsal position. Cover it with sufficient solution to prevent the muscles from drying out.</li>
	<li>Identify the Achilles tendon, carefully raise it with forceps, and dissect the muscles away from the bone with iris scissors. Use a stereomicroscope from this step onward of the dissection.</li>
	<li>Identify and separate the whole gastrocnemius muscle, the main bulk at the back of the hindlimb.</li>
	<li>Dissect and discard the connective and fat tissue surrounding the muscle with fine-tip forceps. Transfer the lateral head of the muscle to a new Petri dish with ice-cold Relax solution (see Figure 2A).</li>
	<li>Gently hold the muscle with forceps from one end and carefully separate it into bundles with microscissors. Always manipulate bundles by gently holding them at one end with forceps. 
	NOTE: Bundles are approximately 10 mm long and 2 mm wide.</li>
	<li>Transfer the fiber bundles to a new Petri dish with 2 mL of ice-cold Relax solution. Only select the ones that have a smooth appearance, are complete from one end to the other, and are not shortened (Figure 2B).</li>
</ol>
3. Live-cell imaging acquisition of mitochondria in skeletal muscle by confocal microscopy
<ol>
	<li>Incubate the fibers in 2.5 &#215; 10-4 mM TMRE in Relax solution for 20 min at room temperature. 
	NOTE: With the recommended working concentrations of TMRE, it is expected to be in a non-quenching mode. Protect from light exposure from this point onwards.</li>
	<li>During the incubation time:
	<ol>
		<li>Open the confocal microscope standard software, select the configuration framework, and in the hardware configuration dialog box select laser and check HeNe 543 option.</li>
		<li>In the acquire framework, select the acquisition dialog box, and in the acquisition mode, select XYZ panel.</li>
		<li>In the XY dialog box, select the 512 x 512 format, 400 Hz speed, and check pinhole panel. In the displayed pinhole dialog box, select AU for unit and add a value of 3 Airy for the pinhole. 
		NOTE: Consider reducing the pinhole size closest to the 1 Airy optimal criterion, if an adequate fluorescence signal is conserved.</li>
		<li>In the Beam Path Settings dialog box, select a water immersion objective of 20x and numerical aperture (NA) of 0.7 (20x/0.7 IMM), and select the emission wavelength window of 576-700 nm. Select DD488/543 and 15% of laser power for the 543 laser. 
		NOTE: A water immersion objective lens for live imaging scanning is strongly recommended (if available). Since a similar refractive index of the immersion medium and the incubation medium is achieved.</li>
	</ol>
	</li>
	<li>After 20 min of incubation, exchange the incubation medium twice. Ensure that the confocal images are acquired within 20-30 min after incubation.</li>
	<li>Prepare the recording chamber with a 0.15 mm coverslip of borosilicate glass. 
	NOTE: Select the thickness of the coverslip according to the available recording chamber, considering that thickness usually ranges from 0.15 to 0.22 mm for optimal performance.</li>
	<li>Add 200 &#181;L of Relax solution to the recording chamber and transfer the fiber bundles.</li>
	<li>In the confocal microscope, use the brightfield mode to identify viable fibers for the fluorescent recording of mitochondria. Viable fibers are complete, not contracted, and have an intact striation pattern.</li>
	<li>Separate the fiber bundles from each other and align them with forceps. Select the ones that are closest to the coverslip.</li>
	<li>Scan the fluorescence using the live button to adjust gain and offset in the Control Panel console considering the following:
	<ol>
		<li>Select low fluorescent intensity values for a background of nearly 0 Arbitrary Units (A.U.).</li>
		<li>Select a gain level between 100 and 200 A.U.&#160;to avoid saturation of the recording system. Hence, do not record the highest fluorescence levels.</li>
		<li>Select a pixel size of 150-190 nm to acquire the full width of the fiber, adjusting the zoom factor in the XY dialog box. 
		NOTE: Consider using the pixel size closer to the Nyquist criteria (90 nm), which allows scanning of the full width of the fiber.</li>
	</ol>
	</li>
	<li>In the Z stack dialog box, adjust the Z distance to acquire the fluorescence signal starting at a fiber depth of 15 &#181;m (begin button) up to 22 &#181;m (end button). Select 3 &#181;m as the Z-step size.</li>
	<li>Click the Start button to acquire the confocal images. Obtain a Z-stack composed of three confocal images acquired at 15, 18, and 21 &#181;m of depth.</li>
</ol>
4. Analysis of mitochondrial density
<ol>
	<li>In the open-source platform for biological image analysis18&#160;open the Z-stack file and rotate the images to place the fiber horizontally, as shown in Figure 3B.</li>
	<li>Randomly select a rectangular region of interest (ROI) that includes the area occupied by the mitochondria (ROImito). Select ROImito sizes ranging from 65 to 90 &#181;m for X. For Y, the size will depend on the fiber width, avoiding its periphery.</li>
	<li>Duplicate the Z-stack with the selected ROImito&#160;and save it as a new Z-stack (Mito-stack) in TIFF format. Save the ROImito position selected on the original stack with the ROI Manager tool.</li>
	<li>Calculate the threshold for background subtraction as follows:
	<ol>
		<li>Use the shortcut command+shift+t to open the Threshold dialog box.</li>
		<li>Select the threshold algorithm Otsu |B&#38;W | Dark background option. Observe that the image is now binarized.</li>
		<li>In the Threshold dialog box, observe the histogram of the fluorescence intensity distribution and the threshold value that is displayed. 
		NOTE: Mitochondria-positive pixels have fluorescence intensity values greater than the threshold and appear as white pixels in the binary image.</li>
		<li>Apply the threshold to the binary image stack by selecting Apply. In the displayed Convert Stack to Binary dialog box, select the options Calculate threshold for each image, Black background, Create new stack, and click OK.</li>
	</ol>
	</li>
	<li>Observe that a stack with the three binary images is generated (BinDMito-stack).&#160;Save it&#160;in TIFF format.</li>
	<li>Calculate mitochondrial density in the BinDMito-stack as follows:
	<ol>
		<li>Select the Analyze menu. Next, click Histogram. In the displayed histogram dialog box, click Yes to include all the images of the stack for analysis.</li>
		<li>In the Histogram of Stack dialog box, click List to obtain the histogram data. Transfer the histogram data to a spreadsheet.</li>
		<li>In a spreadsheet, identify Mito-pixels&#160;that are those with the 255 value. Calculate Mitochondrial density&#160;using equation (1): 
		Mitochondrial density = <img alt="Equation 1" src="/files/ftp_upload/65306/65306eq01.jpg" style="margin: auto;" />&#160;&#215;&#160;100&#160; &#160; &#160;(1) 
		Total pixels are identified as N in the Histogram dialog box or calculated by summing in the spreadsheet the pixels count of the histogram.</li>
		<li>To calculate the area occupied by mitochondria (&#181;m2), multiply the Mito-pixels of the BinDMito-stack&#160;by the pixel size.</li>
	</ol>
	</li>
</ol>
5. Analysis of mitochondrial distribution by Fast Fourier Transform
<ol>
	<li>In the open-source platform for biological image analysis, open the BinDMito-stack&#160;and draw a rectangular ROI of 256 pixels width and 5 &#181;m height. Locate an ROI in a central position and another in a lateral position, as shown in Figure 4A,B.</li>
	<li>Use the ROI Manager tool to configure the ROIs for FFT analysis and save it. 
	NOTE: According to the necessities of the analysis, other ROIs can be selected in different positions, and their sizes can be modified. Nevertheless, the width must equal 2n pixels to perform FFT (e.g., 128, 256, 512 pixels).</li>
	<li>Obtain the plot profile of the ROIs using the shortcut command + k and transfer the data to a spreadsheet.</li>
	<li>In the spreadsheet, fill B and C columns with the data obtained from the plot profile. The B column contains the distance in &#181;m, and the C column the corresponding gray value of the fluorescence intensity (A.U.).</li>
	<li>The D column corresponds to the FFT frequency (event/&#181;m), which will be filled as follows:
	<ol>
		<li>Calculate the sampling frequency (Fs): Fs = 1/&#916;d, where &#916;d is the distance step (the second value of B).</li>
		<li>Calculate the &#916;Fs: &#916;Fs = Fs/N, where N is the number of data points of&#160;B&#160;(256).</li>
		<li>Calculate the stop value of FFT frequency (S): S = (N/2) &#215;&#916;Fs.</li>
		<li>Fill D column as follows:
		<ol>
			<li>The first value of D column equals 0.</li>
			<li>Select the second cell in D column, go to the home menu, select fill, and click series. Select columns. For the step value, use the calculated &#916; Fs. For the stop value, use the calculated S.</li>
		</ol>
		</li>
		<li>Next, fill the E column with the FFT complex values as follows:
		<ol>
			<li>Insert 0 in the first cell of the C column, because the distance 0 must coincide with a signal 0 for the calculation of the FFT.</li>
			<li>Then, go to the Data menu, click Data Analysis, and select Fourier Analysis.</li>
			<li>In the new window, select the range of data points of the fluorescent signal described in column C for the input range. 
			NOTE: The number of data points must be 2n (e.g., 256 or 128 data points of fluorescent signal).</li>
			<li>Select the corresponding range of the E for the output range.</li>
			<li>Select Ok and let the FFT complex values be filled automatically.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Fill F column with the FFT magnitude as follows:
	<ol>
		<li>Use the IMABS function to return the absolute value in F from the complex number in E and multiply by 2/N to normalize (equation (2)): 
		FFT magnitude = (IM.ABS (E1&#8230;En) &#215; (2/N)&#160; &#160; &#160;(2)</li>
	</ol>
	</li>
	<li>Plot the FFT spectrum using the FFT magnitude in F, as a function of FFT frequency in D, until S. Find the point of the maximum peak and its corresponding FFT frequency.</li>
	<li>Convert the FFT frequency of the maximum peak to distance with equation (3). This calculated distance represents the longitudinal distance of the mitochondrial distribution. 
	Distance (&#181;m) = 1/FFT frequency&#160; &#160; &#160;(3) 
	NOTE: Due to FFT symmetry, do not plot the FFT frequency beyond the S value, which corresponds to half of the data points, avoiding duplication of the FFT spectrum.</li>
</ol>
6. Optional preprocessing steps to reduce the image noise before image analysis
<ol>
	<li>Apply a median filter or a 2D deconvolution as follows:
	<ol>
		<li>For median filter:
		<ol>
			<li>Select Process | Filters and click on the Median option.</li>
			<li>In the displayed Median dialog box, select 2.0 for Radius and click OK. Choose Yes to apply the process to all images in the Mito-stack.</li>
			<li>Observe the reduction of noise in the images.</li>
		</ol>
		</li>
		<li>For 2D deconvolution:
		<ol>
			<li>Generate a theoretical Point Spread Function (PSF).</li>
			<li>Download the plugin PSF_Generator.jar and place the file into the &#34;Plugins&#34; folder.</li>
			<li>In the PSF generator dialog box, click the option Born &#38; Wolf 3D optical model, enter the Refractive index immersion value of 1.33 used during confocal scanning, and select Best for the Accuracy computation option.</li>
			<li>Capture 576 nm for emission wavelength, the NA of the objective lens used of 0.7, and the Z-step of 3,000 nm.</li>
			<li>Capture the pixel size XY of the confocal image to be deconvoluted, as well as the size XYZ indicating the number of pixels of X and Y and the number of Z steps (3 for this protocol).</li>
			<li>In the Display menu of the PSF generator, click the option linear, select 8 bits resolution, and then select the grays option for the lookup table (LUT).</li>
			<li>Run the PSF generator and save the theoretical PSF created in TIFF format.</li>
			<li>Make a sum of slices of the PSF created by opening the Z Project tool of the option Stacks that is located in the Image menu.</li>
			<li>In the displayed ZProjection dialog box, select Sum slices for the Projection type, and click OK. Save the new PSF in TIFF format. 
			NOTE: A PSF obtained experimentally by confocal scanning fluorescent microbeads with a known diameter can be used instead of the theoretical PSF.</li>
			<li>Download the plugin &#34;DeconvolutionLab_2.jar&#34;19 and place it into the plugin folder.</li>
			<li>Click the menu Plugins. Next, click DeconvolutionLab2.</li>
			<li>Separate the confocal images from the Mito-stack&#160;using the tool Images to stack from the Stacks option located in the Image menu and save it in TIFF format.</li>
			<li>In the DeconvolutionLab2 dialog box, select one of the images obtained from the Mito-stack. Select the theoretical PSF created, and select the algorithm Richardson-Lucy with 15 iterations.</li>
			<li>Run DeconvolutionLab2, verify signal improvement and noise reduction in the image, and save it in TIFF format.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Continue with step 4.4 for image analysis.</li>
	<li>For the thresholding of the deconvoluted images, convert them to an 8-bit mask during the thresholding process. Create a stack with the binary and deconvoluted images by selecting Images to Stack in the Stacks tool of the Image menu.</li>
	<li>For the FFT analysis of deconvoluted images, the plot profile contains distance in pixel units. Transform the pixel units to &#181;m as follows:
	<ol>
		<li>In the spreadsheet, after completing step 5.4 transfer the data from B to A. Next, fill B with the distance in &#181;m by multiplying each pixel number from A by the pixel size.</li>
	</ol>
	</li>
</ol>

Confocal Microscopic Analysis of Rat Muscle Mitochondria

<ol>
	<li>Lackner, L. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Shaping+the+dynamic+mitochondrial+network.">Shaping the dynamic mitochondrial network.</a> BMC Biology. 12, 35 (2014).</li><li>De Mario, A., Gherardi, G., Rizzuto, R., Mammucari, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skeletal+muscle+mitochondria+in+health+and+disease.">Skeletal muscle mitochondria in health and disease.</a> Cell Calcium. 94, 102357 (2021).</li><li>Chang, C. R., Blackstone, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Dynamic+regulation+of+mitochondrial+fission+through+modification+of+the+dynamin-related+protein+Drp1.">Dynamic regulation of mitochondrial fission through modification of the dynamin-related protein Drp1.</a> Annals of the New York Academy of Sciences. 1201, 34-39 (2010).</li><li>Willingham, T. B., Ajayi, P. T., Glancy, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Subcellular+specialization+of+mitochondrial+form+and+function+in+skeletal+muscle+cells.">Subcellular specialization of mitochondrial form and function in skeletal muscle cells.</a> Frontiers in Cell and Developmental Biology. 9, 757305 (2021).</li><li>Kuznetsov, A. V., 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=Mitochondrial+subpopulations+and+heterogeneity+revealed+by+confocal+imaging:+possible+physiological+role.">Mitochondrial subpopulations and heterogeneity revealed by confocal imaging: possible physiological role.</a> Biochimica et Biophysica Acta. 1757 (5-6), 686-691 (2006).</li><li>D&#237;az-Vegas, A. 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=Mitochondrial+calcium+increase+induced+by+RyR1+and+IP3R+channel+activation+after+membrane+depolarization+regulates+skeletal+muscle+metabolism.">Mitochondrial calcium increase induced by RyR1 and IP3R channel activation after membrane depolarization regulates skeletal muscle metabolism.</a> Frontiers in Physiology. 9, 791 (2018).</li><li>Boncompagni, S., 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=Mitochondria+are+linked+to+calcium+stores+in+striated+muscle+by+developmentally+regulated+tethering+structures.">Mitochondria are linked to calcium stores in striated muscle by developmentally regulated tethering structures.</a> Molecular Biology of the Cell. 20 (3), 1058-1067 (2009).</li><li>Reggiani, C., Marcucci, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+controversial+issue:+Can+mitochondria+modulate+cytosolic+calcium+and+contraction+of+skeletal+muscle+fibers.">A controversial issue: Can mitochondria modulate cytosolic calcium and contraction of skeletal muscle fibers.</a> The Journal of General Physiology. 154 (9), e202213167 (2022).</li><li>Heinzel, F. 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=Remodeling+of+T-tubules+and+reduced+synchrony+of+Ca2++release+in+myocytes+from+chronically+ischemic+myocardium.">Remodeling of T-tubules and reduced synchrony of Ca2+ release in myocytes from chronically ischemic myocardium.</a> Circulation Research. 102 (3), 338-346 (2008).</li><li>Al-Qusairi, L., Laporte, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=T-tubule+biogenesis+and+triad+formation+in+skeletal+muscle+and+implication+in+human+diseases.">T-tubule biogenesis and triad formation in skeletal muscle and implication in human diseases.</a> Skeletal Muscle. 1 (1), 26 (2011).</li><li>P&#233;rez-Trevi&#241;o, P., P&#233;rez-Trevi&#241;o, J., Borja-Villa, C., Garc&#237;a, N., Altamirano, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Changes+in+T-tubules+and+sarcoplasmic+reticulum+in+ventricular+myocytes+in+early+cardiac+hypertrophy+in+a+pressure+overload+rat+model.">Changes in T-tubules and sarcoplasmic reticulum in ventricular myocytes in early cardiac hypertrophy in a pressure overload rat model.</a> Cellular Physiology and Biochemistry. 37 (4), 1329-1344 (2015).</li><li>Celestino-Montes, A., P&#233;rez-Trevi&#241;o, P., Sandoval-Herrera, M. D., G&#243;mez-V&#237;quez, N. L., Altamirano, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relative+role+of+T-tubules+disruption+and+decreased+SERCA2+on+contractile+dynamics+of+isolated+rat+ventricular+myocytes.">Relative role of T-tubules disruption and decreased SERCA2 on contractile dynamics of isolated rat ventricular myocytes.</a> Life Sciences. 264, 118700 (2021).</li><li>Song, L. S., 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=Orphaned+ryanodine+receptors+in+the+failing+heart.">Orphaned ryanodine receptors in the failing heart.</a> Proceedings of the National Academy of Sciences of the United States of America. 103 (11), 4305-4310 (2006).</li><li>Houzelle, A., 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=Human+skeletal+muscle+mitochondrial+dynamics+in+relation+to+oxidative+capacity+and+insulin+sensitivity.">Human skeletal muscle mitochondrial dynamics in relation to oxidative capacity and insulin sensitivity.</a> Diabetologia. 64 (2), 424-436 (2021).</li><li>Call, J. A., 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=Ulk1-mediated+autophagy+plays+an+essential+role+in+mitochondrial+remodeling+and+functional+regeneration+of+skeletal+muscle.">Ulk1-mediated autophagy plays an essential role in mitochondrial remodeling and functional regeneration of skeletal muscle.</a> American Journal of Physiology. Cell Physiology. 312 (6), C724-C732 (2017).</li><li>Fealy, C. E., Grevendonk, L., Hoeks, J., Hesselink, M. K. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skeletal+muscle+mitochondrial+network+dynamics+in+metabolic+disorders+and+aging.">Skeletal muscle mitochondrial network dynamics in metabolic disorders and aging.</a> Trends in Molecular Medicine. 27 (11), 1033-1044 (2021).</li><li>Rivera-Alvarez, I., 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=A+single+session+of+physical+activity+restores+the+mitochondrial+organization+disrupted+by+obesity+in+skeletal+muscle+fibers.">A single session of physical activity restores the mitochondrial organization disrupted by obesity in skeletal muscle fibers.</a> Life Sciences. 256, 117965 (2020).</li><li>Schindelin, J., 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=Fiji:+an+open-source+platform+for+biological-image+analysis.">Fiji: an open-source platform for biological-image analysis.</a> Nature Methods. 9 (7), 676-682 (2022).</li><li>Sage, 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=DeconvolutionLab2:+An+open-source+software+for+deconvolution+microscopy.">DeconvolutionLab2: An open-source software for deconvolution microscopy.</a> Methods. 115, 28-41 (2017).</li><li>Perry, S. W., Norman, J. P., Barbieri, J., Brown, E. B., Gelbard, H. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+membrane+potential+probes,+and+the+proton+gradient:+a+practical+usage+guide.">Mitochondrial membrane potential probes, and the proton gradient: a practical usage guide.</a> Biotechniques. 50 (2), 98-115 (2011).</li><li>Ma, 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=Mitophagy,+mitochondrial+homeostasis,+and+cell+fate.">Mitophagy, mitochondrial homeostasis, and cell fate.</a> Frontiers in Cell and Developmental Biology. 8, 467 (2020).</li><li>Meinild, L. A. -. 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=Exercise+training+increases+skeletal+muscle+mitochondrial+volume+density+by+enlargement+of+existing+mitochondria+and+not+de+novo+biogenesis.">Exercise training increases skeletal muscle mitochondrial volume density by enlargement of existing mitochondria and not de novo biogenesis.</a> Acta Physiologica. 222 (1), (2018).</li><li>Memme, J. M., Erlich, A. T., Phukan, G., Hood, D. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Exercise+and+mitochondrial+health.">Exercise and mitochondrial health.</a> Journal of Physiology. 599 (3), 803-817 (2021).</li><li>Gan, Z., Fu, T., Kelly, D. P., Vega, R. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Skeletal+muscle+mitochondrial+remodeling+in+exercise+and+diseases.">Skeletal muscle mitochondrial remodeling in exercise and diseases.</a> Cell Research. 28 (10), 969-980 (2018).</li><li>Schiaffino, S., Hanzl&#237;kov&#225;, V., Pierobon, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Relations+between+structure+and+function+in+rat+skeletal+muscle+fibers.">Relations between structure and function in rat skeletal muscle fibers.</a> Journal of Cell Biology. 47 (1), 107-119 (1970).</li><li>Vincent, A. E., 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+spectrum+of+mitochondrial+ultrastructural+defects+in+mitochondria+myopathy.">The spectrum of mitochondrial ultrastructural defects in mitochondria myopathy.</a> Scientific Reports. 6, 30610 (2016).</li><li>Mishra, P., Varuzhanyan, G., Pham, A. H., Chan, D. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mitochondrial+dynamics+is+a+distinguishing+feature+of+skeletal+muscle+fiber+types+and+regulates+organellar+compartmentalization.">Mitochondrial dynamics is a distinguishing feature of skeletal muscle fiber types and regulates organellar compartmentalization.</a> Cell Metabolism. 22 (6), 1033-1044 (2015).</li><li>Cheng, A. J., Westerblad, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mechanical+isolation,+and+measurement+of+force+and+myoplasmic+free+[Ca2+]+in+fully+intact+single+skeletal+muscle+fibers.">Mechanical isolation, and measurement of force and myoplasmic free [Ca2+] in fully intact single skeletal muscle fibers.</a> Nature Protocols. 12 (9), 1763-1776 (2017).</li><li>Kuznetsov, A. V., Javadov, S., Margreiter, R., Hagenbuchner, J., Ausserlechner, M. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Analysis+of+mitochondrial+function,+structure,+and+intracellular+organization+in+situ+in+cardiomyocytes+and+skeletal+Muscles.">Analysis of mitochondrial function, structure, and intracellular organization in situ in cardiomyocytes and skeletal Muscles.</a> International Journal of Molecular Sciences. 23 (4), 2252 (2022).</li><li>Pawley, J. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fundamental+limits+in+confocal+microscopy.">Fundamental limits in confocal microscopy.</a> In: Pawley, J. (eds) Handbook of Biological Confocal Microscopy. , (2006).</li><li>Otsu, N. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Threshold+selection+method+from+gray-level+histogram.">Threshold selection method from gray-level histogram.</a> IEEE Transactions on System Man Cybernetics. 9, 62-66 (1979).</li><li>Schermelleh, L., Heintzmann, R., Leonhardt, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+guide+to+super-resolution+fluorescence+microscopy.">A guide to super-resolution fluorescence microscopy.</a> The Journal of Cell Biology. 190 (2), 165-175 (2010).</li><li>Jakobs, S., Stephan, T., Ilgen, P., Br&#252;ser, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Light+microscopy+of+mitochondria+at+the+nanoscale.">Light microscopy of mitochondria at the nanoscale.</a> Annual Review of Biophysics. 49, 289-308 (2020).</li></ol>

The authors declare that there are no conflicts of interest.

Mitochondria are organelles with a high remodeling capacity. Their content, density, and distribution can be rapidly modified through the activation of the mitochondrial fusion and fission mechanisms, known as mitochondrial dynamics1, and the balance between the mitochondrial turnover mechanisms: the mitochondrial biogenesis and the specialized mitochondria degradation pathway, mitophagy21,22. Mitochondrial content and morphology can vary according to cell type and stage of development and can be remodeled under different physiological and pathological stimuli17,22,23,24. Therefore, the study of mitochondrial morphology has been relevant for over half a century25. Notably, the analysis of mitochondria through electron microscopy has been the standard technique applied in multiple studies26.
Fluorescent studies by confocal microscopy have gained relevance in the last few years due to their capacity for live-cell imaging of mitochondria at different fiber depths, which could help better understand the role of mitochondria in skeletal muscle in different adaptive and maladaptive conditions27. In this study, a methodology for the analysis of mitochondria density and distribution in live-skeletal muscle fibers by confocal microscopy is described. One of the main challenges of working with live-skeletal muscle fibers is to avoid contraction from the isolation processes up to the mitochondrial confocal recording. To achieve this goal, a high Mg and ATP Relax solution17 is used to keep the fibers relaxed for at least 2 h, which gives enough time to carry out the fiber isolation process, the fluorophore load, and the acquisition of mitochondrial signal by confocal microscopy. A critical point of the protocol is obtaining the fibers mechanically since it requires high precision and fresh tissue; however, it is possible to obtain viable fiber bundles from rat muscle with this previously used and reported technique28. Obtaining intact fibers allows for preserving the sarcolemma and the intracellular environment, keeping the metabolic and functional crosstalk between cell structures28,29.
Unlike working with tissues or fixed cells, the acquisition of live-cell imaging fluorescent images by confocal microscopy allows real-time monitoring of the effect of diverse experimental conditions. The present protocol can be used to explore changes in mitochondrial density and distribution in real-time and explore differences between experimental groups, such as the examples presented here between Lean and Ob-derived fibers (Figure 3 and Figure 4). It should always be considered that live-cell imaging implies standardizing optimal working conditions with minor cell damage. The working time, the quality of the solutions used, the acquisition parameters, and the exposure of lasers must be finely controlled. Hence, essential considerations are mentioned below.
Mitochondria of muscle fibers cannot be recorded entirely longitudinally by confocal microscopy due to the size of the fiber and the damage of the fiber that can be caused by long laser exposure. Nevertheless, a representative sample of the fiber is recorded under this technique. Although it is possible to record the entire thickness of the skeletal muscle fiber of a rat by confocal microscopy, this implies a longer recording time and exposure to the laser beam. In the case of control rats, issues with these recordings have not been encountered. However, fibers from pathological conditions may be more susceptible to damage as observed in the fibers from Ob rats. Consequently, acquiring a stack of representative confocal images obtained at different Z distances is preferred. When only a section of fiber thickness is recorded, it is recommended to take the stack at the same depth on all fibers tested since mitochondrial distribution and density can vary according to its position within the fiber. Acquisition of the signal at a depth above 15 &#181;m is recommended to obtain representative confocal images of IMF, avoiding SSM populations that are situated close to the periphery.
During the confocal acquisition, some important considerations must be taken into account. First, the selection of the immersion objective lens considering the magnification, high NA, and immersion medium. Since cells are maintained in a hydrophilic incubation medium, the refractive index of the incubation and immersion medium must be similar to obtain a good signal and scan deep into the tissue. Usually achieved using a water immersion objective lens. Confocal images of Figure 3 and Figure&#160;4 were acquired with a 20x, 0.7 NA, water immersion objective. This objective allows the record of the fiber in all its depth, but scanning at 15, 18, and 21 &#181;m was decided since representative confocal images of IMF can be obtained with high fluorescence intensity signal and minor fiber damage. Other magnification, such as 40x and oil as an immersion medium, can be considered but needs to be evaluated.
Second, the pixel size for imaging acquisition is calculated according to the Nyquist theorem, which allows the selection of an appropriate pixel size that avoids over-sample (higher laser exposure) and under-sample (leads to less resolution)30. The calculation depends on the characteristics of the objective lens selected and the wavelength (~90 nm). It can be adjusted with the zoom; therefore, only one zoom setting provides an optimal pixel size30. Nevertheless, in practice, the zoom also depends on the area of the specimen to be analyzed. Thus, finding balance allows working with a pixel size that is closest to the Nyquist criterion and that also fits the area to be analyzed. Figure&#160;3 and Figure&#160;4 were acquired with a pixel size of 150 and 190 nm, which allowed for analysis of the full width of the fiber which is ~50-80 &#181;m.
Third, an appropriate pinhole diameter that prevents out-of-focus light from reaching the detector should be used. Typically, 1 Airy is considered the optimal pinhole size since it allows the detection of ~80% of photons originating from the plane of focus30. Nevertheless, some stained biological samples that show low fluorescence levels require a pinhole increase30. Confocal images of Figure&#160;3 and Figure&#160;4 were acquired with a pinhole size of 3 Airy due to a low signal captured with a lower Airy. It is important to consider that the rise in the signal intensity resulting from increasing the pinhole size leads to the reduction of the resolution due to increased out-of-focus captured light. For this reason, we recommended using a pinhole size as close to 1 airy as possible.
When adequately acquired, confocal images can be processed to obtain quantitative information on mitochondrial density and distribution. Regardless, the critical processing image step of thresholding needs to be performed before analysis to improve the quantification of the signal. During this crucial step, the fluorescence intensity value that separates the positive pixels for mitochondria from those of the background is defined. The threshold can be defined by a Gaussian fit of the peak representing mitochondria when the histogram of the image displays two peaks, one corresponding to the background and the other to the mitochondria. However, a bimodal distribution is not always achieved in each of the images, so other thresholding methods have to be applied.
In this protocol, the implementation of Otsu&#39;s thresholding is described, which is a non-parametric and unsupervised method designed to find the threshold value when the two peaks are not separated, or other peaks are present31. Otsu can easily be applied using an open-source platform for biological-image analysis; however, other thresholding methods can be tested. The same thresholding method must be applied to all the confocal images and has to be calculated independently for each confocal image. Applying the threshold to a whole stack leads to incorrect results. Once the binary images are obtained after the thresholding process, the analysis of mitochondrial density and FFT can be easily carried out by following the instructions described in this protocol. However, care should be taken when performing both analyses to avoid including nuclei and capillaries, as it would lead to quantification errors. Regarding density, it is enough to subtract the pixels, or the area occupied by the nuclei or capillaries, from the pixels or total area to be analyzed. In addition, when performing the FFT analysis, it must be verified that the mitochondria signal is straight. Conversely, when the mitochondria signal is tilted, it can produce profiles that do not represent the mitochondrial longitudinal distribution, yielding incorrect FFT spectrum data. In addition, a preprocessing step can be applied to reduce the noise in the images. This protocol describes two optional preprocessing steps using a median filter and 2D deconvolution. The effects of these preprocessing methods on the image and mitochondrial density content are presented in Supplementary Figure S1. It is important to consider that while these preprocesses can improve the image quality, they can also result in the loss of certain image details. Therefore, they should be used with caution and consistently applied to all the images being analyzed.
Despite its advantages, confocal microscopy is limited by a lateral resolution (XY) of 180-250 nm when the optimal conditions for acquisition are implemented32. Mitochondrial diameter is ~200-700 nm, close to the diffraction limit of confocal microscopy; thus, sub-mitochondrial structures cannot be adequately detected33 and cannot be evaluated by density and FFT analyses shown in this protocol. Other super-resolution techniques of microscopy, such as stochastic optical reconstruction microscopy (STORM), stimulated emission depletion (STED) nanoscopy, or structured illumination microscopy (SIM), can be explored to resolve sub-mitochondrial structures32. In this protocol, the confocal images of mitochondria are obtained using the fluorophore TMRE, which depends on the mitochondrial membrane potential. Therefore, mitochondrial fluorescent intensity can vary according to their membrane potential. A thresholding process is performed before the data analysis to overcome this issue. All the pixels above a defined threshold are considered positive for mitochondrial signal independent of their fluorescence value. Nevertheless, it must be noted that mitochondria with a very low membrane potential cannot be resolved with this technique. Thus, complementary studies of mitochondrial protein content quantification are recommended. An advantage of using TMRE is that confocal images can also be used for mitochondrial membrane potential analysis, but adequate controls with uncoupling agents need to be performed such as carbonylcyanide-p-trifluoromethoxyphenylhydrazone (FCCP). In addition, the instructions for mitochondrial density and distribution analysis can be achieved using green fluorescent indicators for mitochondria, which load mitochondria regardless of their membrane potential, but incubation strategy and confocal acquisition settings need to be standardized.
Given that mitochondria structure is related to essential mitochondrial and cellular functions, the protocol described here can provide valuable information about their remodeling during disease or by a particular stress insult. It could contribute to a better understanding of key functions of skeletal muscle governed by mitochondria, such as energy production, or in which mitochondria play an important role in interacting with other organelles, such as contraction-metabolism coupling. Following the protocol instructions allows the estimation of mitochondrial density and distribution in live-skeletal muscle. The protocol steps are divided into three main stages focused on skeletal muscle bundle dissection, confocal microscopy scanning, and image analysis, where detailed instructions and important considerations are included. Notably, the protocol can be further optimized to explore additional Z steps for full mitochondrial reconstruction within the fiber according to the user&#39;s necessities. For instance, the confocal image and analysis steps can be tested to study cellular structures with similar distribution, such as TT in live and fixed samples.

Mitochondria form highly dynamic networks that are mainly regulated by the balance between its elongation (fusion) and fragmentation (fission)1,2, which are modulated by the expression and activity of proteins such as Mitofusin 1 and 2 (Mfn1 and Mfn2), and Optic protein atrophy 1 (Opa1), which regulate the fusion of the outer mitochondrial membrane and inner membrane, respectively1,2. Dynamin-related protein (Drp1) predominantly regulates mitochondrial fission when it is phosphorylated in the Ser6163.
In skeletal muscle, it has been well established that the mitochondrial network is arranged in structurally well-defined subpopulations based on their proximity to different cell regions (myofibrils, sarcolemma, and nuclei)4,5. Those mitochondria located right beneath the sarcolemma are called subsarcolemmal mitochondria (SSM), those located between the contractile filaments are called intermyofibrillar mitochondria (IMF), and the mitochondrial subpopulation around the nuclei are called perinuclear mitochondria network (PMN). Moreover, it has been suggested that these mitochondrial subpopulations have region-specific functions and are metabolically specialized4,5.
The maintenance of cellular energy homeostasis, which allows metabolic and contractile function, depends to a large extent on the interaction and communication on specific sites through the mitochondrial network (e.g., IMF and SSM interaction)4,6. In addition to mitochondria network interactions, the mitochondrion can also interact with other organelles, forming structural and functional complexes. In this regard, it has been shown that IMF can be located adjacent to the sarcoplasmic reticulum (SR) and in proximity to the Ca2+ release units (CRU), formed by the transverse tubules (TT)7. This fact is relevant due to the role of mitochondrial Ca2+ uptake in regulating ATP synthesis and apoptosis. Recently, a potential role in regulating cytosolic Ca2+ transients has also been suggested8.
TT are invaginations of sarcolemma that have a periodic distribution along the longitudinal axis of cardiomyocytes and skeletal muscle fibers9,10, similar to the IMF distribution5. Changes in the distribution of TT have important physiological implications, given their role in contractile function. However, these changes have been mainly evaluated in cardiomyocytes. Using Fast Fourier Transform (FFT) analysis allows the conversion of periodic signals from the distance domain to the frequency domain, resulting in an FFT spectrum that indicates the frequency and the regularity of the signal11,12,13. Although there is evidence that the organization of the mitochondrial network in skeletal muscle fibers is essential for adaptation to different metabolic conditions, as during regeneration after muscle injury14,15, most analyses are performed qualitatively.
Additionally, given that mitochondrial dysfunction has been associated with several skeletal muscle-related (e.g., disuse atrophy)2 and non-muscle diseases, particularly metabolic diseases, and the associated loss of muscle mass (i.e., atrophy)16, the quantitative evaluation of the mitochondrial network and distribution in skeletal muscle takes relevance. Recently, a significant difference in the longitudinal distribution of mitochondria of gastrocnemius muscle fibers between an obese group (Ob; Zucker fa/fa rats), and a lean group (Lean; Zucker +/+ rats) was identified through FFT17. This study demonstrated the usefulness of the FFT in analyzing the mitochondrial distribution. Therefore, this protocol presents a methodology to study mitochondria in live-skeletal muscle fibers from images obtained by fluorescence confocal microscopy. Mitochondrial density is quantified by background thresholding, and the analysis of longitudinal mitochondrial distribution by FFT analysis is also described. A workflow scheme is presented in Figure 1.

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			<td>Adenosine 5&#8217;-triphosphate disodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>A6419</td>
			<td></td>
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			<td>Borosilicate glass coverslip</td>
			<td>Warner Instruments</td>
			<td>64-0709</td>
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			<td>Calcium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>C5670</td>
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			<td>Confocal microscope</td>
			<td>Leica</td>
			<td>TCS SP5</td>
			<td></td>
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			<td>Confocal microscope software Leica Application Suite&#160;</td>
			<td>Leica</td>
			<td>2.7.3.9723</td>
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			<td>Creatine Phosphokinase</td>
			<td>Sigma-Aldrich</td>
			<td>C3755</td>
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			<td>DeconvolutionLab2 (DeconvolutionLab_2.jar)</td>
			<td>Biomedical Imaging Group, EPFL</td>
			<td>http://bigwww.epfl.ch/deconvolution/deconvolutionlab2/</td>
			<td></td>
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			<td>Dimethyl Sulfoxide</td>
			<td>Sigma-Aldrich</td>
			<td>D2650</td>
			<td></td>
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			<td>DL-Aspartic acid potassium sat hemihydrate</td>
			<td>Sigma-Aldrich</td>
			<td>11240</td>
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			<td>Ethylene glycol-bis(2-aminoethylether)-N,N,N&#180;N&#180;-tetraacetic acid</td>
			<td>Sigma-Aldrich</td>
			<td>E4378</td>
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			<td>Forceps</td>
			<td>Miltex</td>
			<td>MH-18</td>
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			<td>HC PL APO 20x/ 0.7 IMM objective&#160;</td>
			<td>Leica</td>
			<td>506517</td>
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			<td>HEPES</td>
			<td>Sigma-Aldrich</td>
			<td>H3375</td>
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			<td>Iris scissors</td>
			<td>Miltex</td>
			<td>5-304</td>
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			<td>L-(-)-Malic acid</td>
			<td>Sigma-Aldrich</td>
			<td>M7397</td>
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			<td>L-glutamic acid monosodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>G1626</td>
			<td></td>
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			<td>Magnesium chloride hexahydrate</td>
			<td>Sigma-Aldrich</td>
			<td>M2393</td>
			<td></td>
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			<td>Maxchelator</td>
			<td>UC Davis Health</td>
			<td>https://somapp.ucdmc.ucdavis.edu/pharmacology/bers/maxchelator/downloads.htm&#160;</td>
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			<td>Micro scissors</td>
			<td>Miltex</td>
			<td>18-1633</td>
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			<td>Open-source platform for biological-image analysis Fiji&#160;</td>
			<td>Public, maintained by Eliceiri/LOCI group, Jug group, and Tomancak lab.Fiji</td>
			<td>https://fiji.sc/</td>
			<td></td>
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			<td>Phosphocreatine disodium salt hydrate</td>
			<td>Sigma-Aldrich</td>
			<td>P7936</td>
			<td></td>
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			<td>Potassium chloride</td>
			<td>Sigma-Aldrich</td>
			<td>P9333</td>
			<td></td>
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			<td>PSF Generator (PSF_Generator.jar)</td>
			<td>Biomedical Imaging Group, EPFL</td>
			<td>http://bigwww.epfl.ch/algorithms/psfgenerator/&#160;</td>
			<td></td>
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			<td>Recording chamber</td>
			<td>Warner Instruments</td>
			<td>RC-27N</td>
			<td></td>
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			<td>Sodium hydroxide</td>
			<td>Sigma-Aldrich</td>
			<td>S5881</td>
			<td></td>
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			<td>Spreadsheet Microsoft Excel</td>
			<td>Microsoft&#160;</td>
			<td></td>
			<td></td>
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			<td>Stereo microscope</td>
			<td>Zeiss</td>
			<td>Stemi 508</td>
			<td></td>
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			<td>Tetramethylrhodamine, ethyl ester</td>
			<td>Invitrogen</td>
			<td>T669</td>
			<td></td>
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analysis of the mitochondrial density and longitudinal distribution in rat live-skeletal muscle fibers by confocal microscopy

The mitochondrion is an organelle that can be elongated, fragmented, and renovated according to the metabolic requirements of the cells. The remodeling of the mitochondrial network allows healthy mitochondria to meet cellular demands; however, the loss of this capacity has been related to the development or progression of different pathologies. In skeletal muscle, mitochondrial density and distribution changes are observed in physiological and pathological conditions such as exercise, aging, and obesity, among others. Therefore, the study of the mitochondrial network may provide a better understanding of mechanisms related to those conditions.
Here, a protocol for mitochondria imaging of live-skeletal muscle fibers from rats is described. Fibers are manually dissected in a relaxing solution and incubated with a fluorescent live-cell imaging indicator of mitochondria (tetramethylrhodamine ethyl ester, TMRE). The mitochondria signal is recorded by confocal microscopy using the XYZ scan mode to obtain confocal images of the intermyofibrillar mitochondrial (IMF) network. After that, the confocal images are processed by thresholding and binarization. The binarized confocal image accounts for the positive pixels for mitochondria, which are then counted to obtain the mitochondrial density. The mitochondrial network in skeletal muscle is characterized by a high density of IMF population, which has a periodic longitudinal distribution similar to that of T-tubules (TT).&#160;The Fast Fourier Transform (FFT) is a standard analysis technique performed to evaluate the distribution of TT that allows finding the distribution frequency and the level of their organization. In this protocol, the implementation of the FFT algorithm is described for the analysis of the longitudinal mitochondrial distribution in skeletal muscle.

Following the present protocol, the analysis of the density and distribution of mitochondria can be achieved in live-skeletal muscle. The protocol is divided into three main stages: Skeletal muscle bundle dissection, confocal microscopy scanning, and image analysis. The workflow overview is presented in Figure 1. Figure 2A shows a whole rat gastrocnemius muscle in a Petri dish, marking the lateral head from which the fibers are obtained, while Figure 2B shows the fiber bundles in Relax solution. By confocal microscopy, the mitochondria can be recorded along the depth of live-skeletal muscle fiber using the fluorescent indicator TMRE. TMRE is a lipophilic cationic fluorophore that is selectively accumulated within mitochondria according to the mitochondrial membrane potential20.
An optimal confocal image of IMF can be obtained by selecting a Z distance above 15 &#181;m of depth within the fiber (Figure 3A). Figure 3B shows representative XY confocal images of mitochondria loaded with TMRE acquired along gastrocnemius muscle fibers&#39; Z distance (15 to 21 &#181;m). The confocal images were processed by thresholding to transform them into binary images to allow mitochondrial analysis. Figure 3B (left panel)&#160;shows a fiber from an exercised Lean rat. It represents an expected confocal record of skeletal muscle fiber mitochondria since it has a consistent pattern along the fiber. In contrast, we selected a fiber from an Ob rat (Figure 3B, right panel) that shows substantial alterations in mitochondrial content and distribution. Figure 3C shows the quantification of the fiber area occupied by the mitochondria expressed as mitochondrial density, obtained from each binarized image (shown in panel B). As expected, the Ob fiber presented a lower mitochondrial density. It was quantified consistently along the Z distance analyzed, which is also observed in Figure 3D when the mitochondrial density is calculated per stack conformed by the three confocal images acquired at 15, 18, and 21 &#181;m.
Similar to the mitochondrial density analysis, the confocal scanning of a fluorescent indicator for live-cell imaging, such as TMRE, allows for studying the longitudinal organization of mitochondria in live-skeletal muscle. IMF presents a periodic organization in the I-band close to TT7, which can be analyzed by FFT to quantify the frequency of the mitochondrial signal and the level of organization17. Figure 4 shows the differences in the organization of IMF found in gastrocnemius derived from Lean and Ob rats and how FFT allows finding the changes in the distribution of the mitochondrial signal. Figure 4A,B exhibit longitudinal ROIs selected in a central and lateral position in the fiber for FFT analysis. Before performing FFT, the thresholding for background subtraction is calculated. Then, the image is binarized; these procedures eliminate the variations in fluorescence intensity levels of the mitochondrial signal. The binary image provides the plot profile of fluorescence distribution necessary to perform the FFT.
Figure 4C,D show the selected ROIs in panels A and B with their plot profiles (upper panels). From the plot profiles, differences in the fluorescence distribution between the fibers derived from Lean and Ob rats can be observed, as well as the variations between ROIs within the same fiber. For every plot profile, their respective FFT spectrum is presented (lower panels). The peak of the maximum FFT spectrum indicates the FFT frequency (X-axis) of the mitochondrial signal distribution along the longitudinal axis. It can be transformed into a distance value close to 2 &#181;m in the lateral and central ROIs from the Lean rat. Notably, the FFT magnitude of the peak is an index of the regularity of the mitochondrial signal, and changes in this amplitude reveal alterations in the mitochondrial distribution.
Figure 4E,F show the differences in the FFT spectrum between Lean and Ob-derived fibers where lateral and central ROIs were analyzed, respectively. In lateral ROIs (Figure 4E) the frequency of mitochondrial longitudinal distribution was similar in the lean and Ob-derived fibers; nonetheless, the amplitude of the maximum FFT peak in the Ob-derived fibers was higher, which is in agreement with the higher regularity of the signal observed in the image in Figure 4B. Nevertheless, the central ROI (Figure 4F) of the Ob is observed as an example of a critical reduction of the FFT peak compared to Lean when an important alteration of the mitochondrial distribution is present.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65306/65306fig01v2.jpg" /> 
Figure 1: Scheme for mitochondrial analysis in skeletal muscle by confocal microscopy. This scheme summarizes the protocol&#39;s main steps in three separate phases. The first phase, dissection of gastrocnemius muscle fiber bundles, is subdivided into three subsequent steps, gastrocnemius muscle isolation, followed by muscle mechanical dissection into bundles to finally make a visual selection of the viable ones. The second phase consists of live-cell imaging acquisition by confocal microscopy, which consists of incubation with the fluorophore (TMRE) for 20 min at room temperature to place the fibers in the chamber. Afterward, the appropriate settings are made in the microscope to conduct the acquisition of the confocal images. During the third phase, confocal image processing and data analysis are conducted. Starting with image processing, where a threshold is required to generate binary images from which mitochondrial density calculations and mitochondrial distribution by FFT are done. Abbreviations: TMRE = tetramethylrhodamine ethyl ester; FFT = Fast Fourier Transform. <a href="https://www.jove.com/files/ftp_upload/65306/65306fig01largev2.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/65306/65306fig02.jpg" /> 
Figure 2: Skeletal muscle bundle dissection. (A) Dissected&#160;rat lateral gastrocnemius head (black arrow) in a Petri dish with Relax solution. (B) Representative image of gastrocnemius muscle fiber bundles in Relax solution before TMRE loading. Abbreviation: TMRE = tetramethylrhodamine ethyl ester. <a href="https://www.jove.com/files/ftp_upload/65306/65306fig02large.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/65306/65306fig03.jpg" /> 
Figure 3: Mitochondrial density analysis. (A) Illustration that represents the Z distance recommended for confocal scanning of IMF mitochondria in skeletal muscle fibers. (B) Series of confocal images of mitochondria loaded with TMRE in skeletal muscle fibers, obtained from an exercised Zucker +/+ rat (Lean) and from an obese Zucker fa/fa rat (Ob) recorded in the Z distance (from 15 to 21 &#181;m of depth). The images were processed by thresholding and transformed into binary images. (C) Calculated mito-density of the confocal slices obtained at different Z distances observed in panel B. (D) The mito-density obtained from the stack composed by the images observed in panel B. Images in panel B&#160;are 65 x 50&#160;&#181;m. Scale bar = 10 &#181;m. Abbreviations: Mito-density = Mitochondrial density; IMF = intermyofibrillar mitochondria; Ob = Obese; SSM = subsarcolemmal mitochondria. <a href="https://www.jove.com/files/ftp_upload/65306/65306fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65306/65306fig04.jpg" /> 
Figure 4: Mitochondrial distribution analysis by FFT. Representative confocal images of mitochondria loaded with the fluorescent indicator TMRE acquired at 21 &#181;m of the depth of skeletal muscle fibers obtained from an exercised Zucker +/+ rat (Lean, panel A) and from an obese Zucker fa/fa rat (Ob, panel B). The images were processed by thresholding and transformed into binary images. Lateral and central ROIs from panel A (panel C) and panel B (panel D) with their respective plot profiles of fluorescence intensity (upper plot) and FFT spectrum (lower plot). FFT was calculated from ROIs with 256 pixels of width, which corresponds to an ROI size of 39 x 5&#160;&#181;m in panel A and an ROI size of 50 x 5&#160;&#181;m in panel B. Panel E shows the differences of the FFT spectrum found between mitochondria derived from Lean and Ob rats in the lateral ROI, while panel F shows the differences of the FFT spectrum of central ROI. Scale bar = 10 &#181;m. Abbreviations: A.U. = Arbitrary Units; FFT = Fast Fourier Transform; ROIs = regions of interest; Ob = Obese. <a href="https://www.jove.com/files/ftp_upload/65306/65306fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Reagent</td>
			<td>Final Concentration in 100 mL</td>
			<td>Stock&#160;</td>
		</tr>
		<tr>
			<td>K-aspartate</td>
			<td>100 mM</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>20 mM</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>20 mM</td>
			<td></td>
		</tr>
		<tr>
			<td>L-glutamic acid</td>
			<td>3 mM</td>
			<td></td>
		</tr>
		<tr>
			<td>Malic acid</td>
			<td>3 mM</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>0.1 mM</td>
			<td>10 mM</td>
		</tr>
		<tr>
			<td>MgCl2</td>
			<td>1 mM free Mg2+</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2</td>
			<td>0.00002 mM free Ca2+</td>
			<td></td>
		</tr>
		<tr>
			<td>Phosphocreatine di-Na</td>
			<td>5 mM</td>
			<td>500 mM</td>
		</tr>
		<tr>
			<td>Creatine phosphokinase</td>
			<td>5 U/mL</td>
			<td>200 U/mL</td>
		</tr>
		<tr>
			<td>MgATP</td>
			<td>5 mM</td>
			<td></td>
		</tr>
		<tr>
			<td>pH 7.3 (with NaOH)</td>
			<td></td>
			<td></td>
		</tr>
	</tbody>
</table>
Table 1: Relax solution reagents and concentration.
Supplementary Figure S1: Effect of image preprocessing methods. (A) Representative confocal images of mitochondria loaded with the fluorescent indicator TMRE acquired at a depth of 18 &#181;m in skeletal muscle fibers obtained from an exercised Zucker +/+ rat (Lean, upper panel) and from an obese Zucker fa/fa rat (lower panel), along with their respective images obtained after preprocessing using Otsu&#180; thresholding, median filter and Otsu&#180; thresholding, and 2D deconvolution and Otsu&#180; thresholding. (B)&#160;Plot profile of mito-density obtained from the confocal images (panel A) after preprocessing with different methods. Images in panel A are 65 x 50&#160;&#181;m. Scale bar = 10 &#181;m. Abbreviations: 2D-Decon = 2D deconvolution; Mito-density = Mitochondrial density. <a href="https://www.jove.com/files/ftp_upload/65306/Supl 1-rev4.pdf" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Mitochondrial Remodeling in Skeletal Muscle at JoVE.com

Author Spotlight: Mitochondrial Remodeling in Skeletal Muscle 

Here, we present a protocol to analyze changes in mitochondrial density and longitudinal distribution by live-skeletal muscle imaging using confocal microscopy for mitochondrial network scanning.

Analysis of the Mitochondrial Density and Longitudinal Distribution in Rat Live-Skeletal Muscle Fibers by Confocal Microscopy

The mitochondrion is an organelle that can be elongated, fragmented, and renovated according to the metabolic requirements of the cells. The remodeling of ...

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3.0K Views.  Tecnologico de Monterrey. Here, we present a protocol to analyze changes in mitochondrial density and longitudinal distribution by live-skeletal muscle imaging using confocal microscopy for mitochondrial network scanning.

Video: Author Spotlight: Mitochondrial Remodeling in Skeletal Muscle 

In vivo Micro-circulation Measurement in Skeletal Muscle by Intra-vital Microscopy

BACKGROUND: Regulatory factors and detailed physiology of in vivo microcirculation have remained not fully clarified after many different modalities of imaging had invented. While many macroscopic parameters of blood flow reflect flow velocity, changes in blood flow velocity and red blood cell (RBC) flux does not hold linear relationship in the microscopic observations. There are reports of discrepancy between RBC velocity and RBC flux, RBC flux and plasma flow volume, and of spatial and temporal heterogeneity of flow regulation in the peripheral tissues in microscopic observations, a scientific basis for the requirement of more detailed studies in microcirculatory regulation using intravital microscopy. METHODS: We modified Jeff Lichtman's method of in vivo microscopic observation of mouse sternomastoid muscles. Mice are anesthetized,
ventilated, and injected with PKH26L-fluorescently labeled RBCs for microscopic observation.RESULT &amp; CONCLUSIONS: Fluorescently labeled RBCs are detected and distinguished well by a wide-field microscope. Muscle contraction evoked by electrical stimulation induced increase in RBC flux. Quantification of other parameters including RBC velocity and capillary density were feasible. Mice tolerated well the surgery, injection of stained RBCs, microscopic observation, and electrical stimulation. No muscle or blood vessel damage was observed, suggesting that our method is relatively less invasive and suited for long-term observations.

A new versatile method for observation of microcirculation is presented. It is considered suitable for long-term observation, and for combination with pharmacophysiological or molecular biological interventions.

BACKGROUND: Regulatory factors and detailed physiology of in vivo microcirculation have remained not fully clarified after many different modalities of ...

Live Imaging of the Zebrafish Embryonic Brain by Confocal Microscopy

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the developing zebrafish brain (Gutzman et al, 2008). Such analysis affords a new understanding of the underlying cell biology required for development of the 3D structure of the vertebrate brain, and significantly increases our ability to study neural tube morphogenesis. The embryonic zebrafish brain is shaped beginning at 18 hours post fertilization (hpf) as the ventricles within the neuroepithelium inflate. By 24 hpf, the initial steps of neural tube morphogenesis are complete. Using the method described here, embryos at the one cell stage are injected with mRNA encoding membrane-targeted green fluorescent protein (memGFP). After injection and incubation, the embryo, now between 18 and 24 hpf, is mounted, inverted, in agarose and imaged by confocal microscopy. Notably, the zebrafish embryo is transparent making it an ideal system for fluorescent imaging. While our analyses have focused on the midbrain-hindbrain boundary and the hindbrain, this method could be extended for analysis of any region in the zebrafish to a depth of 
80-100 &mu;m.

In this video, we demonstrate a method by which to analyze the developing vertebrate brain in live zebrafish embryos at single cell resolution by confocal microscopy. This includes the method by which we inject the single-cell zebrafish embryo and subsequently mount and image the developing brain.

In this video, we demonstrate the method our lab has developed to analyze the cell shape changes and rearrangements required to bend and fold the ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Intravital Microscopy of the Microcirculation in the Mouse Cremaster Muscle for the Analysis of Peripheral Stem Cell Migration

In the era of intravascular cell application protocols in the context of regenerative cell therapy, the underlying mechanisms of stem cell migration to nonmarrow tissue have not been completely clarified. We describe here the technique of intravital microscopy applied to the mouse cremaster microcirculation for analysis of peripheral bone marrow stem cell migration in vivo. Intravital microscopy of the M. cremaster has been previously introduced in the field of inflammatory research for direct observation of leucocyte interaction with the vascular endothelium. Since sufficient peripheral stem and progenitor cell migration includes similar initial steps of rolling along and firm adhesion at the endothelial lining it is conceivable to apply the M. cremaster model for the observation and quantification of the interaction of intravasculary administered stem cells with the endothelium. As various chemical components can be selectively applied to the target tissue by simple superfusion techniques, it is possible to establish essential microenvironmental preconditions, for initial stem cell recruitment to take place in a living organism outside the bone marrow.

Intravital microscopy of the mouse M. cremaster microcirculation offers a unique and well-standardized in vivo model for the analysis of peripheral bone marrow stem cell migration.

In the era of intravascular cell application protocols in the context of regenerative cell therapy, the underlying mechanisms of stem cell migration to ...

Assessment of Calcium Sparks in Intact Skeletal Muscle Fibers

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in the striated muscle fibers that appear as highly localized Ca2+ release events mediated by ryanodine receptor (RyR) Ca2+ release channels on the sarcoplasmic reticulum (SR) membrane. Proper assessment of muscle Ca2+ sparks could provide information on the intracellular Ca2+&#160;handling properties of healthy and diseased striated muscles. Although Ca2+ sparks events are commonly seen in resting cardiomyocytes, they are rarely observed in resting skeletal muscle fibers; thus there is a need for methods to generate and analyze sparks in skeletal muscle fibers.
Detailed here is an experimental protocol for measuring Ca2+ sparks in isolated flexor digitorm brevis (FDB) muscle fibers using fluorescent Ca2+ indictors and laser scanning confocal microscopy. In this approach, isolated FDB fibers are exposed to transient hypoosmotic stress followed by a return to isotonic physiological solution. Under these conditions, a robust Ca2+ sparks response is detected adjacent to the sarcolemmal membrane in young healthy FDB muscle fibers. Altered Ca2+ sparks response is detected in dystrophic or aged skeletal muscle fibers. This approach has recently demonstrated that membrane-delimited signaling involving cross-talk between inositol (1,4,5)-triphosphate receptor (IP3R) and RyR contributes to Ca2+ spark activation in skeletal muscle. In summary, our studies using osmotic stress induced Ca2+ sparks showed that this intracellular response reflects a muscle signaling mechanism in physiology and aging/disease states, including mouse models of muscle dystrophy (mdx mice) or amyotrophic lateral sclerosis (ALS model).

Described here is a method to directly measure calcium sparks, the elementary units of Ca2+ release from sarcoplasmic reticulum&#160;in intact skeletal muscle fibers. This method utilizes osmotic-stress-mediated triggering of Ca2+ release from ryanodine receptor in isolated muscle fibers. The dynamics and homeostatic capacity of intracellular Ca2+ signaling can be employed to assess muscle function in health and disease.

Maintaining homeostatic Ca2+ signaling is a fundamental physiological process in living cells. Ca2+ sparks are the elementary units of Ca2+ signaling in ...

Analyzing Craniofacial Morphogenesis in Zebrafish Using 4D Confocal Microscopy

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical clarity and amenability to genetic manipulation, the zebrafish embryo has become a popular model organism with which to perform time-lapse analysis of morphogenesis in living embryos. Confocal imaging of a live zebrafish embryo requires that a tissue of interest is persistently labeled with a fluorescent marker, such as a transgene or injected dye. The process demands that the embryo is anesthetized and held in place in such a way that healthy development proceeds normally. Parameters for imaging must be set to account for three-dimensional growth and to balance the demands of resolving individual cells while getting quick snapshots of development. Our results demonstrate the ability to perform long-term in vivo imaging of fluorescence-labeled zebrafish embryos and to detect varied tissue behaviors in the cranial neural crest that cause craniofacial abnormalities. Developmental delays caused by anesthesia and mounting are minimal, and embryos are unharmed by the process. Time-lapse imaged embryos can be returned to liquid medium and subsequently imaged or fixed at later points in development. With an increasing abundance of transgenic zebrafish lines and well-characterized fate mapping and transplantation techniques, imaging any desired tissue is possible. As such, time-lapse in vivo imaging combines powerfully with zebrafish genetic methods, including analyses of mutant and microinjected embryos.

Time-lapse confocal imaging is a powerful technique useful for characterizing embryonic development. Here, we describe the methodology and characterize craniofacial morphogenesis in wild-type, as well as pdgfra, smad5, and smo mutant embryos.

Time-lapse imaging is a technique that allows for the direct observation of the process of morphogenesis, or the generation of shape. Due to their optical ...

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and temporal resolution to study in vivo hematopoiesis using the murine bone marrow transplant experimental model. Genetic combinatorial marking using lentiviral vectors encoding fluorescent proteins (FPs) enabled cell fate mapping through advanced microscopy imaging. Vectors encoding five different FPs: Cerulean, EGFP, Venus, tdTomato, and mCherry were used to concurrently transduce HSPCs, creating a diverse palette of color marked cells. Imaging using confocal/two-photon hybrid microscopy enables simultaneous high resolution assessment of uniquely marked cells and their progeny in conjunction with structural components of the tissues. Volumetric analyses over large areas reveal that spectrally coded HSPC-derived cells can be detected non-invasively in various intact tissues, including the bone marrow (BM), for extensive periods of time following transplantation. Live studies combining video-rate multiphoton and confocal time-lapse imaging in 4D demonstrate the possibility of dynamic cellular and clonal tracking in a quantitative manner.

In vivo Clonal Tracking of Hematopoietic Stem and Progenitor Cells Marked by Five Fluorescent Proteins using Confocal and Multiphoton Microscopy

Combinatorial 5 fluorescent proteins marking of hematopoietic stem and progenitor cells allows in vivo clonal tracking via confocal and two-photon microscopy, providing insights into bone marrow hematopoietic architecture during regeneration. This method allows non-invasive fate mapping of spectrally-coded HSPCs-derived cells in intact tissues for extensive periods of time following transplantation.

We developed and validated a fluorescent marking methodology for clonal tracking of hematopoietic stem and progenitor cells (HSPCs) with high spatial and ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

Simultaneous Measurement of Mitochondrial Calcium and Mitochondrial Membrane Potential in Live Cells by Fluorescent Microscopy

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ concentration. To do this, mitochondria utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester Ca2+. The influx of Ca2+ into the mitochondria stimulates three rate-limiting dehydrogenases of the citric acid cycle, increasing electron transfer through the oxidative phosphorylation (OXPHOS) complexes. This stimulation maintains &#916;&#936;m, which is temporarily dissipated as the positive calcium ions cross the mitochondrial inner membrane into the mitochondrial matrix.
We describe here a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using confocal microscopy. By permeabilizing the cells, mitochondrial Ca2+ can be measured using the fluorescent Ca2+ indicator Fluo-4, AM, with measurement of &#916;&#936;m using the fluorescent dye tetramethylrhodamine, methyl ester, perchlorate (TMRM). The benefit of this system is that there is very little spectral overlap between the fluorescent dyes, allowing accurate measurement of mitochondrial Ca2+ and &#916;&#936;m simultaneously. Using the sequential addition of Ca2+ aliquots, mitochondrial Ca2+ uptake can be monitored, and the concentration at which Ca2+ induces mitochondrial membrane permeability transition and the loss of &#916;&#936;m determined.

Mitochondria can utilize the electrochemical potential across their inner membrane (&#916;&#936;m) to sequester calcium (Ca2+), allowing them to shape cytosolic Ca2+ signaling within the cell. We describe a method for simultaneously measuring mitochondria Ca2+ uptake and &#916;&#936;m in live cells using fluorescent dyes and confocal microscopy.

Apart from their essential role in generating ATP, mitochondria also act as local calcium (Ca2+) buffers to tightly regulate intracellular Ca2+ ...

A Rapid Automated Protocol for Muscle Fiber Population Analysis in Rat Muscle Cross Sections Using Myosin Heavy Chain Immunohistochemistry

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle composition. Various time-consuming methods have traditionally been used to study fiber populations in many fields of research. However, recently developed immunohistochemical methods based on myosin heavy chain protein expression provide a quick alternative to identify multiple fiber types in a single section. Here, we present a rapid, reliable and reproducible protocol for improved staining quality, allowing automatic acquisition of whole cross sections and automatic quantification of fiber populations with ImageJ. For this purpose, embedded skeletal muscles are cut in cross sections, stained using myosin heavy chains antibodies with secondary fluorescent antibodies and DAPI for cell nuclei staining. Whole cross sections are then scanned automatically using a slide scanner to obtain high-resolution composite pictures of the entire specimen. Fiber population analyses are subsequently performed to quantify slow, intermediate and fast fibers using an automated macro for ImageJ. We have previously shown that this method can identify fiber populations reliably to a degree of &#177;4%. In addition, this method reduces inter-user variability and time per analyses significantly using the open source platform ImageJ.

Here, we present a protocol for rapid muscle fiber analyses, which allows improved staining quality, and thereby automatic acquisition and quantification of fiber populations using the freely available software ImageJ.

Quantification of muscle fiber populations provides a deeper insight into the effects of disease, trauma, and various other influences on skeletal muscle ...