Three-dimensional Imaging and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

Skin punch biopsies were obtained from subjects that were recruited from a large community-based primary care network at the University of Utah Diabetes Center (Salt Lake City, UT). This study was approved by the University of Michigan Institutional Review Board and complied with the tenets of the Declaration of Helsinki. Written informed consent was obtained from each subject prior to testing.

1. Fluorescence Immunohistochemistry

1. Prepare punch biopsies for intraepidermal nerve fiber immunohistochemistry:
   1. Perform 3 mm skin biopsies by medical staff and place the whole biopsy in 1.5 mL Zamboni's fixative solution (2% paraformaldehyde, 0.3% saturated picric acid in phosphate buffered saline (PBS), pH 7.4) at 4 °C overnight.
   2. Rinse samples with a solution of 30% sucrose in PBS at 4 °C for 16-24 h or until the sample sinks.
   3. Embed samples in optimal cutting temperature compound (OCT) using a cryomold. Place the whole 3 mm biopsy with the epidermis facing down in the mold and fill the mold with approximately 2 mL of OCT. Freeze the mold on crushed dry ice. Store at -80 °C until ready to use.
   4. Cut 50 µm thick cross sections using a cryostat and store in individual wells of a 96-well plate using 180 µL antifreeze storage solution per well (30% ethylene glycol, 30% glycerol in PBS). The following directions are for 8 wells of a 96 well plate. Stain sections from each biopsy site that are 200 – 300 µm away from each other.

DAY 1:

2. Quench non-specific labeling of the stratum corneum:
   1. Label 96-well plate as shown in Figure 1.
   2. Pipet 150 µL of stock signal enhancer solution to reduce nonspecific binding of secondary antibodies^34,35 into each well. Transfer sections into signal enhancer solution using an inoculating loop.
      NOTE: Take care while working with the tissue to avoid damaging or tearing the tissue sections. Keep in signal enhancer solution for 30 minutes at room temperature on a flat rocker.
   3. Prepare rinse wells in rows 2 and 3 of the 96-well plate by adding 150 µL of 1x phosphate buffered saline (PBS) into each well.
   4. Carefully transfer sections into row 2 and rinse in 1x PBS for 10 min at room temperature.
   5. Rinse a second time in 1x PBS for 10 min at room temperature (row 3).
3. Prepare 5% bovine serum albumin (BSA) blocking solution:
   1. Prepare 5% blocking solution that contains 5% BSA and 0.3% Triton X 100 (TX-100) in 0.1 M PBS (see Table 1) while sections are incubating in signal enhancer solution. BSA does not go into solution easily. Vortex solution until BSA completely dissolves.
   2. Prepare blocking wells in row 4 of the 96-well plate by adding 150 µL of 5% BSA blocking solution into each well.
   3. Transfer sections into individual wells of 5% BSA blocking solution and incubate sections in 5% BSA blocking solution for 1 – 2 h at room temperature on a flat rocker.
4. Prepare 1% BSA rinsing solution and dilution of primary antibodies:
   1. Prepare 1% rinsing solution that contains 1% BSA and 0.3% TX-100 in 0.1 M PBS (see Table 2) while sections are incubating in 5% BSA blocking solution. BSA does not go into solution easily. Vortex solution until BSA completely dissolves.
   2. Dilute primary antibodies in 1% BSA rinsing solution while sections are incubating in 5% BSA blocking solution.
      1. Make 1,500 µL of primary antibody solution and add to each well in row 5.
      2. Dilute the primary antibodies: Use the nerve-specific label, rabbit polyclonal anti-protein gene product 9.5 (PGP9.5) at 1:1,000. Use the mitochondria-specific label, mouse monoclonal anti-pyruvate dehydrogenase E2/E3bp antibody (PDH) at 1:100 in 1% BSA rinsing solution.
5. Prepare primary antibody:
   1. Pipet 150 µL of primary antibody into each well in row 5 of the 96-well plate.
   2. Using the loop tool, transfer sections from the blocking solution (row 4) into the row 5 containing the primary antibody.
   3. Wrap the plate tightly with parafilm to keep it from drying out.
   4. Place samples on flat rocker at room temperature for 1 h, and then incubate samples at 4 ºC on a flat rocker overnight.

DAY 2:

6. Rinse samples:
   1. Prepare rinse wells in rows 6, 7 and 8 of the 96-well plate by adding 150 µL of 1% BSA rinsing solution into each well.
   2. Transfer sections into first 1% BSA rinsing solution (row 6) and incubate for 1 h at room temperature. Repeat rinses twice more by incubating sections in 1% BSA rinsing solution in rows 7 and 8 for 1 h each at room temperature.
7. Dilute secondary antibodies in 1% BSA rinsing solution:
   1. Make 1,500 µL of secondary antibody solution while sections are incubating in the last rinse of 1% BSA rinsing solution (row 8).
   2. Dilute secondary antibodies: for PGP9.5 (green-fluorescent conjugated goat anti-rabbit antibody, 1:1000), for PDH (red-fluorescent conjugated goat anti-mouse, 1:1,000) in 1% BSA rinsing solution.
8. Prepare secondary antibody:
   1. Pipet 150 µL of secondary antibody into row 9 of the 96-well plate.
   2. Gently transfer the sections from 1% BSA rinsing solution (row 8) into secondary antibody wells (row 9).
   3. Using parafilm, wrap the plate tightly to keep it from drying out. Cover with aluminum foil. Place samples on flat rocker at room temperature for 1 h, and then incubate samples at 4 ˚C on a flat rocker overnight.

DAY 3:

9. Prepare clean 1x PBS by filtering through a 0.22 µm filter:
   1. Pipet 150 µL of filtered 1x PBS into row 10, 11, and 12.
   2. Transfer samples to row 10 and rinse in 1x PBS for 1 h at room temperature. Cover the 96-well plate with aluminum foil and place on a flat rocker during rinse. Repeat filtered 1x PBS rinse two more times for 1 h each at room temperature in rows 11 and 12.
10. Prepare microscope slides for mounting tissue sections:
    1. Place 50 µL of filtered 1x PBS on a slide.
    2. Transfer section from 1x PBS (row 12) into the 50 µL drop. Carefully position the section in the drop of PBS by unfolding the tissue and gently flattening it onto the glass slide. Remove excess PBS with a glass pipette to avoid diluting the mounting reagent. Do not touch sections with the glass bulb.
    3. Pipet 1 – 2 drops of mounting reagent containing DAPI directly on top of the section on the microscope slide using care to not disturb the orientation of the section. Gently place a 50 mm x 24 mm #1.5 microscope glass coverslip over the section.
    4. Clear any air bubbles that formed while placing the coverslip and wipe excess liquid off the edges of the coverslip. Prepare a new slide and repeat the mounting process for each section.
    5. Cure/dry the mounting media by placing the slides in the dark overnight at room temperature. Transfer the slides to 4 °C for short-term (1 – 2 weeks) or -20 °C for long-term storage (greater than 2 weeks).
       NOTE: Negative controls omitted primary antibodies for PGP9.5 or PDH in step 1.4.2.2 and displayed no distinguishable labeling of nerves or mitochondria in the epidermis. Positive controls were performed for the PDH antibody to prove it labels all mitochondria (data not shown). Positive controls were performed on cultured primary mouse dorsal root ganglion neurons that were transduced with a baculovirus to label mitochondria with a Green Fluorescent Protein (GFP) signal and then fixed and stained with the PDH antibody and red fluorescent secondary antibody. All mitochondria that were expressing GFP were co-labeled with the red label for PDH immunohistochemistry (data not shown).

2. Confocal Imaging

1. Perform confocal imaging:
   1. Collect images using a laser scanning confocal microscope with a 40X oil-immersion (1.25 numerical aperture (N.A.)) objective on an inverted microscope.
      1. At each focal plane sequentially acquire fluorescent signals:
         Nuclei: excitation λ = 405 nm, spectral emission filter λ = 420 – 480 nm
         Nerve fibers: excitation λ= 488 nm, spectral emission filter λ = 505 – 560 nm
         Mitochondria: excitation λ = 543 nm, spectral emission filter λ= 606 – 670 nm
   2. Enter the following scan parameters into the microscope software: scan rate of 600 Hz with 2 frame averaging and zoom of 2.2; 12-bit intensity resolution (4096 gray levels).
      1. Set the microscope software for optimized lateral resolution (scan resolution = 1,024 x 1,024) and axial resolution/optical sectioning (confocal aperture = 1 airy unit (AU) with z-step size of 210 nm).
         NOTE: The resulting XYZ resolution is 172.2 nm x 172.2 nm x 210 nm with an image size of 176.1 µm x 176.1 µm x 30-50 µm.
   3. Activate a live scan for the nerve signal (green-fluorescence) and adjust the z-focus control to find and set the upper and lower focal planes in the microscope software that encompass the nerve signal within the tissue section. The total z-range is typically 30-50 µm for a 50 µm tissue section.
   4. Rotate the scan field with the microscope software during a live scan so that the epidermis is horizontally or vertically positioned in the image.
   5. Scan each signal separately and adjust the detector (photomultiplier tube, PMT) voltage and offset to minimize/remove any over and under saturated pixels.
      NOTE: Scan times with the above parameters take approximately 20 – 40 min depending on the number of z slices.

3. 3D Visualization and Analysis of Mitochondria within Human Intraepidermal Nerve Fibers

1. Isolate the 3D epidermis:
   1. Duplicate the original image and use the maximum intensity projection (extended focus view) of the image to identify and isolate the epidermis.
   2. Use a region of interest tool to trace along the upper and lower edges of the epidermis to remove unwanted areas such as the stratum corneum and dermis that are absent of intraepidermal nerve fibers. Crop to this selection.
2. Use deconvolution on the nerve and mitochondrial fluorescent signals:
   NOTE: Deconvolution helps to restore the integrity of the fluorescent signals. The restoration used in this protocol is called blind deconvolution because it uses the fluorescent signals in the images to determine how much the signals spread from their original source (point spread function). The process improves signal resolution by reassigning the signal spread back to its origin location.
   1. Calculate a point spread function (PSF) for the green fluorescent nerve signal (green-fluorescence) with the following parameters:
      1. Set calculated PSF to confocal. Set medium refractive index to 1.515 and numerical aperture to 1.25. Set detector pinhole to 1 Airy unit (A.U.). Set laser excitation wavelength to 488 nm and emission wavelength to 515 nm.
   2. Calculate a PSF for the red fluorescent mitochondrial signal (red-fluorescence) with the following parameters:
      1. Set calculated PSF to confocal. Set medium refractive index to 1.515 and numerical aperture to 1.25. Set detector pinhole to 1 AU. Set laser excitation wavelength to 543 nm and emission wavelength to 617 nm.
   3. Optimize the nerve and mitochondria fluorescent signals by deconvolution using the corresponding PSFs listed above and iterative restoration feature set at 100% confidence and an iteration limit of 10 cycles.
3. Create nerve-specific surfaces:
   1. Use the "create surface" tool to make a solid surface of the nerves from the deconvolved green- fluorescent secondary labeling of the PGP9.5 identified nerves.
   2. Uncheck the "smooth" feature and use the absolute intensity feature to set the threshold for the nerve signal since it is significantly brighter than background fluorescence.
   3. Use the absolute intensity feature to set the threshold for the nerve signal since it is significantly brighter than background fluorescence. Set threshold value low enough to accurately identify the nerves.
   4. Filter out small, non-nerve surfaces based on size.
      NOTE: If necessary, manually edit out additional non-nerve surfaces within the "Edit" tab by holding the CONTROL key to highlight multiple objects and then deleting them with the Delete key.
4. Isolate nerve-specific fluorescent mitochondrial signal:
   1. Select the Edit tab of the nerve surface to view the "Mask Properties" feature. The nerve surface created in steps 3.3 is used to isolate mitochondria within those nerves away from mitochondrial signals associated with the keratinocytes.
   2. As the "Mask All' button opens a "Mask Channel" window, choose the deconvolved red-fluorescent signal from the pull down menu under "Channel Selection" for the mitochondrial signal.
   3. Click in the box to put a check mark in the "duplicate channel before applying mask" option.
   4. Click on the radio button in front of the "Constant inside/outside" option of the Mask Settings and click in the box to put a check mark in the "Set voxel outside surface to" option and type in 0.00 for the value. Click OK button to create the new channel that represents mitochondrial signals within the nerve surface.
5. Create mitochondria-specific surfaces:
   1. Use the "create surface" tool to make a solid surface of the mitochondria from the newly created fluorescent channel of the nerve-specific mitochondrial signals from steps 3.4.
   2. Uncheck the "smooth" feature and select the "background subtraction" feature to set the threshold. This feature uses local contrast around the mitochondrial signal to identify mitochondria from background.
   3. Set threshold value low enough to accurately identify mitochondria. In this example the lower threshold was set at 2,000 for a 16-bit (65,536) scale.
   4. Filter mitochondrial surfaces based on size. In this example the voxel limit was set to 1.0 voxels, which is the lowest limit possible. If necessary, manually edit out non-mitochondria surfaces within the "Edit" tab by holding the CONTROL key to select multiple objects and then deleting them with the Delete key.
      NOTE: Occasionally, the software will create surfaces that are not associated with a distinguishable fluorescent mitochondrial signal. In these cases, it is possible to remove them with the "Edit" tab.
6. Export and calculate morphometric values for analysis:
   1. Export values for nerve and mitochondrial surfaces from the "Statistics" tab for further analysis with electronic spreadsheet software.
   2. Export the volume values for both the nerve and mitochondrial surfaces.
   3. Count the number of individual nerve fibers present in each image and record the value in the electronic spreadsheet.
   4. For each image, calculate the following potential values for analysis in the electronic spreadsheet:
      1. Sum the volumes for all nerve surfaces.
      2. Filter out nerve-specific mitochondrial surfaces below a volume of less than 0.02 µm³ and bin the surfaces by size. For example, use bins such as 0.02 – 0.04, 0.04 – 0.08, 0.08 – 0.16, 0.16 – 0.32, 0.32 – 0.64, 0.64 – 1.28, 1.28 – 2.56, greater than 2.56 µm³.
         NOTE: The lower limit of mitochondrial volume was set to 0.02 µm³ based on previously published volumes of mitonchodria^36,37,38.
      3. Count the number of nerve-specific mitochondrial surfaces within each bin and the total number of surfaces over the bins (0.02 – greater than 2.56 µm³).
      4. Calculate the proportion of nerve-specific mitochondrial surfaces in each bin. Use the count per bin divided by the total number of mitochondria surfaces.
      5. Sum the volumes for all nerve-specific mitochondrial surfaces.
      6. Calculate the proportion of nerve with mitochondrial signal by dividing the total nerve surface volume by the sum of all mitochondrial surface volumes.
      7. Calculate the number of mitochondria per nerve volume by dividing the total nerve surface volume by the count of all mitochondrial surfaces.