This work was supported by a grant from the National Institutes of Health, National Institute of Diabetes and Digestive and Kidney Diseases, 2R01DK59558 (to G.N.). UAMS Translational Research Institute supported by the National Institutes of Health National Center for Research Resources grant UL1 RR029884 provided partial funding for Flow Cytometry Core at UAMS. We thank Dr. Peipei Ping (University of California at Los Angeles; Los Angeles, CA) for providing an aliquot of adenovirus carrying cDNA coding the dominant negative (inactive) mutant of PKC-&epsilon; and Dr. Allen Samarel (Loyola University Medical Center; Maywood, IL) for providing an aliquot of adenoviral vector coding the constitutively active mutant of PKC-&epsilon;. We also thank Drs. Peter Parker and Peter Sugden (Imperial College London, London, UK) for providing cDNA coding constitutively active PKC-&epsilon;.

1. Isolation of Renal Proximal Tubules for Primary Culture
<ol>
	<li>Anesthetize the rabbit, excise both kidneys and place them in a Petri dish filled with sterile prep buffer (DMEM/F12 medium) in a laminar flow hood.</li>
	<li>Perfuse each kidney with 50 ml of prep buffer followed by 50 ml of sterile phosphate-buffered saline, pH 7.4 (PBS), and PBS-iron oxide solution (45 ml + 5 ml). De-capsulate kidneys and transfer them to the prep buffer containing deferoxamine.</li>
	<li>Harvest the cortex of each kidney.</li>
	<li>Homogenize the tissue using a 15 ml Dounce homogenizer. Pour the homogenate over 2 sterile mesh sieves (85 &mu;m on the bottom and 235 &mu;m on top) placed over a 1 L sterile beaker. Rinse the sieves with deferoxamine-containing buffer.</li>
	<li>Collect any tissue left on the 85 &mu;m sieve in a sterile conical centrifuge tube filled with 35-40 ml of the deferoxamine-containing buffer. Gently mix the cell suspension.</li>
	<li>Place a strong magnet outside of the tube to remove iron filled glomeruli and let the suspension settle. Aspirate iron from the magnet using a Pasteur pipette. Repeat this process.</li>
	<li>Prepare 1 ml of digestion medium containing 2,400-2,500 units of collagenase I and 0.6 mg soybean trypsin inhibitor dissolved in the deferoxamine-containing buffer. Filter-sterilize the digestion medium.</li>
	<li>Adjust the volume of the cell suspension to 40 ml, and add 1 ml of digestion medium. Incubate at room temperature for 17 min gently mixing the suspension, centrifuge at 50 x g for 2 min at 4 &deg;C, aspirate the medium, and add fresh deferoxamine buffer.</li>
	<li>Repeat the iron-removing procedure with the magnet several times, spin the suspension down again, aspirate medium, and add 46 ml of medium containing no glucose.</li>
	<li>Mix gently and measure out 500 &mu;l of the suspension into two pre-weighed Eppendorf tubes. Spin cells down at 10,000 x g for 1 min, aspirate the media, and weigh the pellet. Calculate the total yield of wet mass and protein.</li>
	<li>Plate the cells in sterile 35 mm culture dishes at the density of 1 mg protein/dish using glucose-free DMEM/F12 medium. Culture cells in 2 ml glucose-free DMEM/F12 medium always on an orbital shaker in the incubator at 37 &deg;C (95% air/5% CO2).1,2</li>
</ol>
2. Renal Proximal Tubule Cell Culture
<ol>
	<li>The culture medium is a 50:50 mixture of DMEM and Ham's F-12 nutrient mix without phenol red, pyruvate, and glucose, supplemented with 15 mM NaHCO3, 15 mM HEPES, and 6 mM lactate (pH 7.4, 295 mosmol/kgH2O). The media are supplemented with L-ascorbic acid-2-phosphate (50 &mu;M), bovine insulin (10 nM), human transferrin (5 mg/ml), hydrocortisone (50 nM), and selenium (5 ng/ml) immediately before daily media change.</li>
	<li>Aspirate old media from dishes using a sterile technique. Add 2 ml of warm, fresh media to each dish using a sterile technique. Renal proximal tubular cells (RPTC) reach confluence in approximately 5-6 days after plating.1,2</li>
</ol>
3. Adenoviral Infection of RPTC
<ol>
	<li>Adenoviral infection is carried out in confluent, quiescent cultures of RPTC after the daily media change. Dominant negative PKC-&epsilon; mutant (dnPKC-&epsilon;) was constructed by the replacement of: 1) lysine with arginine at position 436 of the ATP-binding site and 2) alanine with glutamate at position 159 of the pseudosubstrate domain. These mutations destroyed the construct's kinase activity without changing the active conformation of PKC-&epsilon;.3 PKC-&epsilon; was made constitutively active by deletion of residues 154-163 of its inhibitory pseudosubstrate domain.4</li>
	<li>Add stock solution of the adenovirus carrying caPKC-&epsilon; cDNA or dnPKC-&epsilon; cDNA to each dish to obtain desired multiplicity of infection (MOI) resulting in significant increase in PKC-&epsilon; protein levels. Incubate RPTC with adenovirus for 24 hr. Change media and culture cells for another 24 hr. Significantly increased levels of phosphorylated and total PKC-&epsilon; proteins can be obtained using 25-50 MOI of adenoviral vector coding caPKC-&epsilon;. Greater MOI results in even larger increases in the levels of active PKC-&epsilon;, but it is not recommended in RPTC due to the rapid cell death and loss. Significantly increased levels of the inactive PKC-&epsilon; protein can be obtained using 50-100 MOI in RPTC without producing adverse effects.5</li>
	<li>At 48 hr after infection, aspirate media, wash the monolayers with PBS, and collect cells for immunoblot analysis to determine protein levels of PKC-&epsilon;.5</li>
</ol>
4. Measurement of RPTC Respiration
<ol>
	<li>Prepare an assay buffer for measuring state 3 respiration (120 mM KCl, 5 mM KH2PO4, 10 mM HEPES, 1 mM MgSO4, and 2 mM EGTA, pH 7.4). For measuring complex I-coupled state 3 respiration, supplement the assay buffer with 5 mM glutamate, 5 mM malate and 0.1 mg/ml digitonin. For complex II-coupled state 3 respiration, supplement the assay buffer with 10 mM succinate, 0.1 &mu;M rotenone, and 0.1 mg/ml digitonin. For complex IV-coupled state 3 respiration, supplement the assay buffer with 1 mM ascorbate, 1 mM N,N,N',N'-tetramethyl-p-phenylenediamine (TMPD), and 0.1 mg/ml digitonin. Place the solutions in a 37 &deg;C water bath.</li>
	<li>Aspirate media from a culture dish containing RPTC monolayer, add 2 ml of a warm state 3 respiration buffer, gently scrape cells from the dish using a rubber policeman, and transfer the suspension to the oxygen consumption chamber equipped with a Clark-type electrode.</li>
	<li>Measure state 2 respiration (in the absence of ADP). Initiate state 3 respiration by adding ADP to a final concentration of 0.4 mM. Initiate state 4 respiration by adding oligomycin to a final concentration of 0.5 &mu;g/ml. Uncoupled respiration is measured by adding 0.5 &mu;M FCCP to cells respiring at state 3.6,7</li>
	<li>Collect the cell suspension from the chamber, precipitate protein using perchloric acid, and spin down the precipitate. Determine protein concentration in the cellular pellet.</li>
</ol>
5. Analysis of Mitochondrial Membrane Potential (&Delta;&Psi;m)
<ol>
	<li>Prepare 0.5 mM JC-1 solution in the sterile culture medium.</li>
	<li>Add slowly 20 &mu;l of JC-1 solution to each dish to obtain final concentration of 5 &mu;M. Swirl the dish to mix the dye thoroughly. Return the dishes to the incubator for 30 min.</li>
	<li>Put dishes on ice, aspirate media, and wash monolayers twice with ice-cold PBS. Aspirate PBS, add 1 ml of fresh PBS, scrape cells from the dish and transfer them to an Eppendorf tube. Break up monolayers into single cells by pipetting them up and down.</li>
	<li>Spin the suspension down at 1,000 x g for 2 min, aspirate the supernatant, add 1 ml of PBS to the pellet and re-suspend it to obtain a single cell suspension. Repeat this step again if necessary.</li>
	<li>Spin cells down at 1,000 x g for 2 min, aspirate the supernatant, add 0.5 ml of PBS, and re-suspend the pellet. Analyze fluorescence by flow cytometry using excitation by a 488-nm argon-ion laser. Read the emission in two separate channels, FL1 at 525 nm and FL2 at 590 nm. Mitochondrial membrane potential is expressed as FL2/FL1 fluorescence ratio.6</li>
</ol>
6. Isolation of Mitochondria
<ol>
	<li>Prepare isolation buffers: Buffer A (10 mM HEPES, 225 mM mannitol, 75 mM sucrose, 0.1 mM EGTA, pH 7.4), Buffer B (buffer A containing protease inhibitors, 1 mM Na3VO4, 1 mM NaF), and Buffer C (10 mM HEPES, 395 mM sucrose, 0.1 mM EGTA, pH 7.4). Perform all steps on ice.</li>
	<li>Wash RPTC monolayers twice with ice-cold PBS buffer. Scrape monolayers into Eppendorf tubes and spin them at 500 x g for 5 min. Wash the pellet in 1 ml of Buffer A.</li>
	<li>Re-suspend the pellet in 1 ml of Buffer B, and transfer the suspension to a 2 ml Dounce homogenizer.</li>
	<li>Homogenize cells in the Dounce homogenizer until most of the cells in the homogenate are broken when inspected under a microscope.</li>
	<li>Centrifuge the homogenate at 1,000 x g for 5 min at 4 &deg;C. Collect the supernatant and spin it down at 15,000 x g for 10 min at 4 &deg;C.</li>
	<li>Discard the supernatant and re-suspend the pellet containing crude mitochondria in 1 ml of Buffer C. Spin the supernatants down at 15,000 x g for 10 min at 4 &deg;C. Repeat the washing of the pellet two more times.5</li>
	<li>To measure activities of respiratory complexes, re-suspend mitochondrial pellet in 50-100 &mu;l of the assay buffer and freeze in liquid nitrogen. Thaw samples slowly on ice.</li>
</ol>
7. Measurement of Activity of NADH-ubiquinone Oxidoreductase (Complex I)
<ol>
	<li>Prepare the assay buffer: 10 mM KH2PO4, 5 mM MgCl2, pH 7.2. Add fatty acid-free BSA and NADH to obtain final concentrations of 0.25% and 0.25 mM, respectively. Add 2 &mu;l of antimycin A solution (1 mg/ml) to a final concentration of 2 &mu;g/ml.</li>
	<li>Run the samples in a 48-well transparent plate. Include a blank sample that has all additions but no mitochondria. To each well, add 500 &mu;l of the assay buffer with additions and mitochondria, and incubate the plate at 30 &deg;C with mixing for 5 min.</li>
	<li>Start the reaction by adding 10 &mu;l of 3.25 mM ubiquinone to each well. Record the absorbance (340 nm) for 3 min in 20 sec intervals. Add 5 &mu;l of rotenone solution (1 mg/ml) to each well and record the absorbance for additional 2 min. Calculate complex I activity as the rotenone-sensitive NADH oxidation.7</li>
</ol>
8. Measurement of Activity of Succinate-ubiquinone Oxidoreductase (Complex II)
<ol>
	<li>Prepare the assay buffer containing 0.25% fatty acid-free BSA, 20 mM potassium succinate, 0.25 mM 2,6-dichloroindophenol, 2 &mu;g/ml antimycin A, and 10 &mu;g/ml rotenone.</li>
	<li>Run the samples in duplicates in a 48-well transparent plate including a blank sample that has all additions but no mitochondria. To each well, add 500 &mu;l of the assay buffer with additions and mitochondria, and incubate the plate at 30 &deg;C with mixing for 2 min.</li>
	<li>Start the reaction by adding 10 &mu;l of 3.25 mM ubiquinone to each well. Record the absorbance (595 nm) for 3 min in 20 sec intervals. Calculate Complex II activity by following the reduction of 2,6-dichloroindophenol. 7</li>
</ol>
9. Measurement of Activity of Ubiquinol-cytochrome c Oxidoreductase (Complex III)
<ol>
	<li>Prepare the assay buffer containing 0.1% fatty acid-free BSA, 80 mM potassium succinate, 10 &mu;g/ml rotenone, and 0.24 mM potassium cyanide.</li>
	<li>Run the samples in duplicates in a 48-well transparent plate including a blank sample that has all additions but no mitochondria. To each well, add 291 &mu;l of the assay buffer with additions and mitochondria, and incubate the plate at 30 &deg;C with mixing for 5 min.</li>
	<li>Start the reaction by adding 3 &mu;l of 6 mM oxidized cytochrome c to each well. Record the absorbance (550 nm) for 3 min in 20 sec intervals. Add 5 &mu;l of antimycin A solution (1 mg/ml) to each well and record the absorbance for additional 2 min. Calculate complex III activity as the antimycin A-sensitive cytochrome c reduction.7</li>
</ol>
10. Measurement of Activity of Cytochrome Oxidase (Complex IV)
<ol>
	<li>Prepare reduced cytochrome c by adding sodium ascorbate to 9 mM solution of cytochrome c and dialyzing the solution overnight at 4 &deg;C in 1 L of phosphate buffer (10 mM KH2PO4, pH 7.2) containing 5 mM MgCl2. Prepare the assay buffer containing 0.1% fatty acid-free BSA and antimycin A (2 &mu;g/ml).</li>
	<li>Run the samples in a 48-well transparent plate including a blank sample that has all additions but no mitochondria. To each well, add 233 &mu;l of the assay buffer with additions and mitochondria, and incubate the plate at 30 &deg;C with mixing for 5 min.</li>
	<li>Start the reaction by addition of reduced cytochrome c (90 &mu;M final concentration). Record the absorbance (550 nm) for 3 min in 20 sec intervals. Add 2 &mu;l of KCN solution (8 &mu;g/ml) to each well and record the absorbance for another 2 min. Calculate complex IV activity as the KCN-sensitive cytochrome c oxidation.7</li>
</ol>
11. Measurement of Activity of F0F1-ATPase (ATP synthase)
<ol>
	<li>Prepare F0F1-ATPase assay buffer (10 mM Tris-HCl, 200 mM KCl, 2 mM MgCl2 , pH 8.2). Re-suspend mitochondrial pellet in the assay buffer, freeze mitochondrial samples in liquid nitrogen and thaw the samples slowly on ice.</li>
	<li>Run the samples in triplicates in a 48-well transparent plate. For each treatment group, 2 wells will be run to measure total ATPase activity (275 &mu;l of the assay buffer, 5 &mu;l mitochondrial suspension, and 5 &mu;l of 95% ethanol) and 1 well to measure oligomycin-insensitive ATPase activity (275 &mu;l of the assay buffer, 5 &mu;l mitochondrial suspension, and 5 &mu;l of 1 mg/ml oligomycin solution in 95% ethanol).</li>
	<li>Incubate samples at 31 &deg;C for 10 min with constant shaking. Add 16 &mu;l of 100 mM ATP solution (pH 7.0) and incubate at 31 &deg;C for 5 min with constant shaking.</li>
	<li>Transfer 200 &mu;l of the incubation mixture to a tube containing 50 &mu;l of 3 M TCA. Incubate samples on ice for 10 min and spin them down at 10,000 x g for 5 min at 4 &deg;C. Use supernatants and pellets to determine phosphate and protein content, respectively.</li>
	<li>Determine phosphate content in the supernatants using the inorganic phosphate assay. Prepare 100 ml of Sumner Reagent by adding 0.88 g FeSO4 to 10 ml of 3.75 M H2SO4 and adjusting to 90 ml with H2O. Add 10 ml of ammonium molybdate solution (0.66 g/10 ml H2O) to the solution of FeSO4 in H2SO4. Protect from light.</li>
	<li>Load 96-well plate with 50 &mu;l of each phosphate standard (0 - 1,000 &mu;M KH2PO4) and 50 &mu;l of each supernatant in duplicates. Add 250 &mu;l of the Sumner Reagent to each well and incubate the plate at 30 &deg;C for 15 min with constant shaking.</li>
	<li>Record the absorbance at 595 nm at room temperature. F0F1-ATPase activity is determined by measuring the oligomycin-sensitive release of Pi from ATP as we described previously.7,8Calculate total and oligomycin-insensitive F0F1-ATPase activities. To calculate the oligomycin-sensitive ATPase activity, subtract oligomycin-insensitive activity from the total ATPase activity. 8,9</li>
</ol>
12. Measurement of Intracellular ATP Content
<ol>
	<li>Place culture dishes containing RPTC monolayers on ice, aspirate medium, and wash the monolayers twice with ice cold PBS buffer. Add 1 ml PBS to each monolayer, scrape each monolayer into PBS, and collect the cell suspension in an Eppendorf tube. Spin the Eppendorf tubes in the microfuge for 5 sec.</li>
	<li>Determine ATP content in each RPTC sample by the luciferase method using the ATP Bioluminescence Assay Kit HS II (Roche) and the manufacturer's protocol. Determine protein concentration in each sample and calculate ATP concentration per mg protein.8</li>
</ol>
13. Visualization of Mitochondrial Morphology
<ol>
	<li>Change culture media. Add 2 &mu;l of 100 &mu;M solution of MitoTracker Red 580 to each dish (final conc. 100 nM) and return the dishes to the incubator for 30 min.</li>
	<li>Examine the live monolayers under a fluorescent microscope using a water immersion objective.5</li>
</ol>
14. Analysis of Cell Viability
<ol>
	<li>At 24 hr before starting the assay prepare 50 mM solution of cisplatin to treat cells that will serve as positive control for apoptosis (annexin V-positive cells). Also, prepare 500 mM solution of tert-butylhydroperoxide to treat cells that will serve as positive controls for oncosis (propidium iodide-positive cells).</li>
	<li>Treat 2 dishes with 2 &mu;l of 50 mM cisplatin and 2 dishes with 2 &mu;l of 500 mM tert-butylhydroperoxide. Dedicate 1 dish for a no-stain control.</li>
	<li>At 24 hr after treatment with cisplatin and TBHP, prepare the binding buffer (10 mM Hepes, 140 mM NaCl, 5 mM KCl, 1 mM MgCl2, 1.8 mM CaCl2) and propidium iodide (PI) solution (50 &mu;g/ml) in the binding buffer.</li>
	<li>Wash monolayers once with the binding buffer. Add 1 ml of the ice-cold buffer and 50 &mu;l of PI solution to each dish except for the no-stain and annexin V-positive cells. Cover dishes and incubate on ice for 15 min with gentle orbital shaking.</li>
	<li>Wash monolayers with the binding buffer (10 min) and add 1 ml of the binding buffer and 2 &mu;l of Annexin V-FITC solution to each dish except for no-stain and PI-positive cells. Cover dishes and incubate at RT for 10 min with gentle orbital shaking.</li>
	<li>Aspirate the buffer and wash the monolayers with 1 ml of the ice-cold binding buffer on ice for 10 min with gentle orbital shaking. Repeat the washing step.</li>
	<li>Aspirate the binding buffer and add 500 &mu;l of binding buffer to each dish. Gently scrape the cells using a rubber policeman and transfer the cells to flow cytometry tubes. Disperse the cells into a single cell suspension by pipetting samples up and down. Break up any cell clusters.</li>
	<li>Quantify PI and annexin V-FITC fluorescence immediately by flow cytometry using excitation at 488 nm and emission at 590 and 530 nm for PI and FITC, respectively. Count 10,000 events for each sample.</li>
	<li>Put the FITC fluorescence on the X-axis and the PI fluorescence on the Y-axis of the flow cytometry dot plot figures. Cells positive for Annexin V and negative for PI are considered apoptotic. Cells positive for PI and negative for Annexin V are considered oncotic.10,11</li>
</ol>

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No conflicts of interest declared.

The approach presented here allows for overexpression of individual isozymes of PKC in the primary culture of renal proximal tubular cells. There are several strengths of this approach: 1. It allows for investigating regulatory mechanisms in a homogenous population of cells (renal proximal tubular cells) that are the primary target for various insults (ischemia, hypoxia, oxidative stress), drugs, and nephrotoxicants within the kidney. 2. Mitochondrial functions in this in vitro model of RPTC grown in pri...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Laminar flow hood</td>
 <td>Thermo Electron Corporation</td>
 <td>FORMA 1104</td>
 </tr>
 <tr>
 <td>2 ml and 15 ml Dounce tissue grinder</td>
 <td>WHEATON</td>
 <td>989-24607, 357544</td>
 </tr>
 <tr>
 <td>85 and 235 micron nylon mesh</td>
 <td>Small Parts</td>
 <td>CMN - 0085 - 10YD CMN - 0250 - 10YD</td>
 </tr>
 <tr>
 <td>50 ml sterile centrifuge tube </td>
 <td>BIOLOGIX</td>
 <td>BCT-P 50BS</td>
 </tr>
 <tr>
 <td>1.5 ml micro tube</td>
 <td>Sarstedt</td>
 <td>72.690.001</td>
 </tr>
 <tr>
 <td>35 x 10 mm sterile culture dishes</td>
 <td>Corning</td>
 <td>430165</td>
 </tr>
 <tr>
 <td>Jouan Centrifuge</td>
 <td>Jouan </td>
 <td>Jouan CR3 11175704 Rotors: Jouan T40</td>
 </tr>
 <tr>
 <td>Adjustable micro-centrifuge</td>
 <td>SIGMA</td>
 <td>Model 1 - 15</td>
 </tr>
 <tr>
 <td>Biological Oxygen Monitor</td>
 <td>YSI Incorporated</td>
 <td>YSI Model 5300A</td>
 </tr>
 <tr>
 <td>Single Chamber Micro Oxygen System</td>
 <td>YSI Incorporated</td>
 <td>5356S</td>
 </tr>
 <tr>
 <td>Oxygen Probe</td>
 <td>YSI Incorporated</td>
 <td>5331A</td>
 </tr>
 <tr>
 <td>Circulating Bath</td>
 <td>YSI Incorporated</td>
 <td>5310</td>
 </tr>
 <tr>
 <td>KCl and Standard Membrane Kit</td>
 <td>YSI Incorporated</td>
 <td>5775</td>
 </tr>
 <tr>
 <td>Magnetic Stirrer</td>
 <td>YSI Incorporated</td>
 <td>5222</td>
 </tr>
 <tr>
 <td>Flatbed Recorder</td>
 <td>Kipp &amp; Zonen</td>
 <td>BD 11E</td>
 </tr>
 <tr>
 <td>48-well and 96-well transparent plates</td>
 <td>Costar</td>
 <td>3548, 3679</td>
 </tr>
 <tr>
 <td>Thermomixer R</td>
 <td>Eppendorf</td>
 <td>5355 21919</td>
 </tr>
 <tr>
 <td>Orbital shaker MAXQ 2000</td>
 <td>Thermo Scientific</td>
 <td>SHKA 2000</td>
 </tr>
 <tr>
 <td>Spectra FLUOR Plus (absorbance/fluorescence/luminescence reader) </td>
 <td>Tecan</td>
 <td>F129005</td>
 </tr>
 <tr>
 <td>Water-Jacketed US Autoflow Automatic CO2 Incubator</td>
 <td>NUAIRE</td>
 <td>NU 4850</td>
 </tr>
 <tr>
 <td>12x75 mm polystyrene culture test tubes for flow cytometry</td>
 <td>Fisher Brand</td>
 <td>14-961-20</td>
 </tr>
 <tr>
 <td>Axioskop 
 Water immersion objective 63x / 0,90W</td>
 <td>Carl Zeiss</td>
 <td>114846 
 ACHROPLAN 44 00 67</td>
 </tr>
 <tr>
 <td>DMEM / F12</td>
 <td>Cellgro</td>
 <td>99 - 830 - PB</td>
 </tr>
 <tr>
 <td>DMEM / F12 Ham</td>
 <td>Sigma</td>
 <td>D 2906 - 1L</td>
 </tr>
 <tr>
 <td>Deferoxamine Mesylate</td>
 <td>Hospira</td>
 <td>D110</td>
 </tr>
 <tr>
 <td>Collagenase Type I</td>
 <td>Worthington</td>
 <td>4196</td>
 </tr>
 <tr>
 <td>Trypsin inhibitor</td>
 <td>Sigma</td>
 <td>T 6522 - 500mg</td>
 </tr>
 <tr>
 <td>5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide (JC-1)</td>
 <td>Invitrogen</td>
 <td>T3168</td>
 </tr>
 <tr>
 <td>Mitotracker Red </td>
 <td>Invitrogen</td>
 <td>M22425</td>
 </tr>
 <tr>
 <td>ATP Bioluminescence Assay Kit HS II</td>
 <td>Roche</td>
 <td>11 699 709 001</td>
 </tr>
 <tr>
 <td>Annexin V - FITC solution</td>
 <td>BioVision</td>
 <td>1001 - 200</td>
 </tr>
 <tr>
 <td>Flow cytometer</td>
 <td>BD Biosciences</td>
 <td>BD FACSCalibur</td>
 </tr>
 </tbody>
</table>

assessment of mitochondrial functions and cell viability in renal cells overexpressing protein kinase c isozymes

The protein kinase C (PKC) family of isozymes is involved in numerous physiological and pathological processes. Our recent data demonstrate that PKC regulates mitochondrial function and cellular energy status. Numerous reports demonstrated that the activation of PKC-a and PKC-&epsilon; improves mitochondrial function in the ischemic heart and mediates cardioprotection. In contrast, we have demonstrated that PKC-&alpha; and PKC-&epsilon; are involved in nephrotoxicant-induced mitochondrial dysfunction and cell death in kidney cells. Therefore, the goal of this study was to develop an in vitro model of renal cells maintaining active mitochondrial functions in which PKC isozymes could be selectively activated or inhibited to determine their role in regulation of oxidative phosphorylation and cell survival. Primary cultures of renal proximal tubular cells (RPTC) were cultured in improved conditions resulting in mitochondrial respiration and activity of mitochondrial enzymes similar to those in RPTC in vivo. Because traditional transfection techniques (Lipofectamine, electroporation) are inefficient in primary cultures and have adverse effects on mitochondrial function, PKC-&epsilon; mutant cDNAs were delivered to RPTC through adenoviral vectors. This approach results in transfection of over 90% cultured RPTC.
Here, we present methods for assessing the role of PKC-&epsilon; in: 1. regulation of mitochondrial morphology and functions associated with ATP synthesis, and 2. survival of RPTC in primary culture. PKC-&epsilon; is activated by overexpressing the constitutively active PKC-&epsilon; mutant. PKC-&epsilon; is inhibited by overexpressing the inactive mutant of PKC-&epsilon;. Mitochondrial function is assessed by examining respiration, integrity of the respiratory chain, activities of respiratory complexes and F0F1-ATPase, ATP production rate, and ATP content. Respiration is assessed in digitonin-permeabilized RPTC as state 3 (maximum respiration in the presence of excess substrates and ADP) and uncoupled respirations. Integrity of the respiratory chain is assessed by measuring activities of all four complexes of the respiratory chain in isolated mitochondria. Capacity of oxidative phosphorylation is evaluated by measuring the mitochondrial membrane potential, ATP production rate, and activity of F0F1-ATPase. Energy status of RPTC is assessed by determining the intracellular ATP content. Mitochondrial morphology in live cells is visualized using MitoTracker Red 580, a fluorescent dye that specifically accumulates in mitochondria, and live monolayers are examined under a fluorescent microscope. RPTC viability is assessed using annexin V/propidium iodide staining followed by flow cytometry to determine apoptosis and oncosis.
These methods allow for a selective activation/inhibition of individual PKC isozymes to assess their role in cellular functions in a variety of physiological and pathological conditions that can be reproduced in in vitro.

Figure 1 shows that adenoviral delivery of cDNA coding the constitutively active (caPKC-&epsilon;) and inactive (dnPKC-&epsilon;) mutants of PKC-&epsilon; results in significantly increased protein levels of PKC-&epsilon; in RPTC and in mitochondria. Cells infected with cDNA carrying the caPKC-&epsilon; vector overexpressed the phosphorylated (active) form of PKC-&epsilon; whereas cells infected with cDNA coding dnPKC-&epsilon; overexpressed PKC-&epsilon; that was inactive (not phosphorylated) (<st...

Watch this Scientific Journal Video about Assessment of Mitochondrial Functions and Cell Viability in Renal Cells Overexpressing Protein Kinase C Isozymes at JoVE.com

Assessment of Mitochondrial Functions and Cell Viability in Renal Cells Overexpressing Protein Kinase C Isozymes

The effects of activation of protein kinase C (PKC) isozymes on mitochondrial functions associated with respiration and oxidative phosphorylation and on cell viability are described. The approach adapts adenoviral technique to selectively overexpress PKC isozymes in primary cell culture and a variety of assays to determine mitochondrial functions and energy status of the cell.

The protein kinase C (PKC) family of isozymes is involved in numerous physiological and pathological processes. Our recent data demonstrate that PKC ...

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18.1K Views.  University of Arkansas for Medical Sciences. The effects of activation of protein kinase C (PKC) isozymes on mitochondrial functions associated with respiration and oxidative phosphorylation and on cell viability are described. The approach adapts adenoviral technique to selectively overexpress PKC isozymes in primary cell culture and a variety of assays to determine mitochondrial functions and energy status of the cell.

Video: Assessment of Mitochondrial Functions and Cell Viability in Renal Cells Overexpressing Protein Kinase C Isozymes

Counting and Determining the Viability of Cultured Cells

Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. A hemacytometer is a thick glass slide with a central area designed as a counting chamber. Cell suspension is applied to a defined area and counted so cell density can be calculated. 

Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. In this video, we demonstrate how cells are counted using a hemacytometer.

Determining the number of cells in culture is important in standardization of culture conditions and in performing accurate quantitation experiments. A ...

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1. Apoptosis can be triggered when cells encounter a wide range of cytotoxic stresses. These insults initiate signaling cascades that ultimately cause the release of cytochrome c from the mitochondrial intermembrane space to the cytoplasm2. The release of cytochrome c from mitochondria is a key event that triggers the rapid activation of caspases, the key cellular proteases which ultimately execute cell death3-4.
The pathway of apoptosis is regulated at points upstream and downstream of cytochrome c release from mitochondria5. In order to study the post-mitochondrial regulation of caspase activation, many investigators have turned to direct cytoplasmic microinjection of holocytochrome c (heme-attached) protein into cells6-9. Cytochrome c is normally localized to the mitochondria where attachment of a heme group is necessary to enable it to activate apoptosis10-11. Therefore, to directly activate caspases, it is necessary to inject the holocytochrome c protein instead of its cDNA, because while the expression of cytochrome c from cDNA constructs will result in mitochondrial targeting and heme attachment, it will be sequestered from cytosolic caspases. Thus, the direct cytosolic microinjection of purified heme-attached cytochrome c protein is a useful tool to mimic mitochondrial cytochrome c release and apoptosis without the use of toxic insults which cause cellular and mitochondrial damage.
In this article, we describe a method for the microinjection of cytochrome c protein into cells, using mouse embryonic fibroblasts (MEFs) and primary sympathetic neurons as examples. While this protocol focuses on the injection of cytochrome c for investigations of apoptosis, the techniques shown here can also be easily adapted for microinjection of other proteins of interest.

Activation of Apoptosis by Cytoplasmic Microinjection of Cytochrome c

In this protocol, we describe the direct cytoplasmic microinjection of cytochrome c protein into fibroblasts and primary sympathetic neurons. This technique allows for the introduction of cytochrome c protein into the cytoplasm of cells and mimics the release of cytochrome c from mitochondria, which occurs during apoptosis.

Apoptosis, or programmed cell death, is a conserved and highly regulated pathway by which cells die1.  Apoptosis can be triggered when cells encounter a ...

Assessment of GFP Expression and Viability Using the Tali Image-Based Cytometer

Single-cell and population information are commonly obtained either by flow cytometry or fluorescence microscopy. However, these two methods provide different information. Flow cytometry gives quantitative multi-parametric information about physical characteristics and staining or expression, but doesn't allow for visualization. Stand-alone fluorescence microscopy provides visual data, but doesn't allow for straightforward quantitative measurements1.
Image-based cytometry bridges the gap between these two methods, enabling the quick visualization and simultaneous quantitative analysis of thousands of cells in heterogeneous populations2. Here, we present a method for performing cell viability and green fluorescent protein (GFP) expression assays using the Tali Image-Based Cytometer3. The Tali instrument is a 3-channel (bright field, green fluorescence, red fluorescence) benchtop assay platform that offers several advantages over flow cytometry and fluorescence microscopy. The Tali cytometer is less expensive, takes up less bench space, requires less maintenance, and the work flow has been simplified so that the operation and analysis is much simpler and quicker. The Tali cytometer is capable of performing a range of suspension cell-based assays, including GFP and red fluorescent protein (RFP) expression, apoptosis4-6 and cell viability analysis with propidium iodide (PI)7-11.
Here, we demonstrate the use of the Tali instrument in performing a cell viability assay in cells expressing GFP. GFP-transduced cells are stained using the Tali Viability Kit - Dead Cell Red. The cells are then pipetted into a Tali Cellular Analysis Slide and loaded into the cytometer. Bright field, red fluorescence and green fluorescence images are captured and analyzed using assay specific algorithms. Histograms are then generated to display cell size, PI fluorescence intensity, and GFP fluorescence intensity. These parameters can then be thresholded to home in on a specific cell population.
A side-by side comparison of the Tali Image-Based Cytometer and traditional flow cytometry demonstrates that the two methods provide comparable data regarding cell viability and protein expression. However, the Tali instrument provides additional visual information about the cell population that cannot be obtained using a flow cytometer.

This protocol describes how to perform cell viability and fluorescence expression assays using the Tali Image-Based Cytometer.

Single-cell and population information are commonly obtained either by flow cytometry or fluorescence microscopy. However, these two methods provide ...

4D Imaging of Protein Aggregation in Live Cells

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental conditions and an intracellular environment that is tightly packed, sticky, and hazardous to protein stability1. The ability to dynamically balance protein production, folding and degradation demands highly-specialized quality control machinery, whose absolute necessity is observed best when it malfunctions. Diseases such as ALS, Alzheimer's, Parkinson's, and certain forms of Cystic Fibrosis have a direct link to protein folding quality control components2, and therefore future therapeutic development requires a basic understanding of underlying processes. Our experimental challenge is to understand how cells integrate damage signals and mount responses that are tailored to diverse circumstances.
The primary reason why protein misfolding represents an existential threat to the cell is the propensity of incorrectly folded proteins to aggregate, thus causing a global perturbation of the crowded and delicate intracellular folding environment1. The folding health, or "proteostasis," of the cellular proteome is maintained, even under the duress of aging, stress and oxidative damage, by the coordinated action of different mechanistic units in an elaborate quality control system3,4. A specialized machinery of molecular chaperones can bind non-native polypeptides and promote their folding into the native state1, target them for degradation by the ubiquitin-proteasome system5, or direct them to protective aggregation inclusions6-9.
In eukaryotes, the cytosolic aggregation quality control load is partitioned between two compartments8-10: the juxtanuclear quality control compartment (JUNQ) and the insoluble protein deposit (IPOD) (Figure 1 - model). Proteins that are ubiquitinated by the protein folding quality control machinery are delivered to the JUNQ, where they are processed for degradation by the proteasome. Misfolded proteins that are not ubiquitinated are diverted to the IPOD, where they are actively aggregated in a protective compartment.
Up until this point, the methodological paradigm of live-cell fluorescence microscopy has largely been to label proteins and track their locations in the cell at specific time-points and usually in two dimensions. As new technologies have begun to grant experimenters unprecedented access to the submicron scale in living cells, the dynamic architecture of the cytosol has come into view as a challenging new frontier for experimental characterization. We present a method for rapidly monitoring the 3D spatial distributions of multiple fluorescently labeled proteins in the yeast cytosol over time. 3D timelapse (4D imaging) is not merely a technical challenge; rather, it also facilitates a dramatic shift in the conceptual framework used to analyze cellular structure.
We utilize a cytosolic folding sensor protein in live yeast to visualize distinct fates for misfolded proteins in cellular aggregation quality control, using rapid 4D fluorescent imaging. The temperature sensitive mutant of the Ubc9 protein10-12 (Ubc9ts) is extremely effective both as a sensor of cellular proteostasis, and a physiological model for tracking aggregation quality control. As with most ts proteins, Ubc9ts is fully folded and functional at permissive temperatures due to active cellular chaperones. Above 30 &deg;C, or when the cell faces misfolding stress, Ubc9ts misfolds and follows the fate of a native globular protein that has been misfolded due to mutation, heat denaturation, or oxidative damage. By fusing it to GFP or other fluorophores, it can be tracked in 3D as it forms Stress Foci, or is directed to JUNQ or IPOD.

Cellular viability depends on timely and efficient management of protein misfolding. Here we describe a method for visualizing the different potential fates of a misfolded protein: refolding, degradation, or sequestration in inclusions. We demonstrate the use of a folding sensor, Ubc9ts, for monitoring proteostasis and aggregation quality control in live cells using 4D microscopy.

One of the key tasks of any living cell is maintaining the proper folding of newly synthesized proteins in the face of ever-changing environmental ...

Viability Assays for Cells in Culture

Manual cell counts on a microscope are a sensitive means of assessing cellular viability but are time-consuming and therefore expensive. Computerized viability assays are expensive in terms of equipment but can be faster and more objective than manual cell counts. The present report describes the use of three such viability assays. Two of these assays are infrared and one is luminescent. Both infrared assays rely on a 16&#160;bit Odyssey Imager. One infrared assay uses the DRAQ5 stain for nuclei combined with the Sapphire stain for cytosol and is visualized in the 700 nm channel. The other infrared assay, an In-Cell Western, uses antibodies against cytoskeletal proteins (&#945;-tubulin or microtubule associated protein 2) and labels them in the 800 nm channel. The third viability assay is a commonly used luminescent assay for ATP, but we use a quarter of the recommended volume to save on cost. These measurements are all linear and correlate with the number of cells plated, but vary in sensitivity. All three assays circumvent time-consuming microscopy and sample the entire well, thereby reducing sampling error. Finally, all of the assays can easily be completed within one day of the end of the experiment, allowing greater numbers of experiments to be performed within short timeframes. However, they all rely on the assumption that cell numbers remain in proportion to signal strength after treatments, an assumption that is sometimes not met, especially for cellular ATP. Furthermore, if cells increase or decrease in size after treatment, this might affect signal strength without affecting cell number. We conclude that all viability assays, including manual counts, suffer from a number of caveats, but that computerized viability assays are well worth the initial investment. Using all three assays together yields a comprehensive view of cellular structure and function.

Therapeutic compounds are often first examined in vitro with viability assays. Blind cell counts by a human observer can be highly sensitive to small changes in cell number but do not assess function. Computerized viability assays, as described here, can assess both structure and function in an objective manner.

Manual cell counts on a microscope are a sensitive means of assessing cellular viability but are time-consuming and therefore expensive. Computerized ...

Experimental Approaches to Study Mitochondrial Localization and Function of a Nuclear Cell Cycle Kinase, Cdk1

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The nuclear-encoded proteins are transported into mitochondria guided by their mitochondria targeting sequences (MTS); however, a majority of mitochondrial localized proteins lack an identifiable MTS. Nevertheless, the fact that MTS can instruct proteins to go into the mitochondria provides a valuable tool for studying mitochondrial functions of normally nuclear and/or cytoplasmic proteins. We have recently identified the cell cycle kinase CyclinB1/Cdk1 complex in the mitochondria. To specifically study the mitochondrial functions of this complex, mitochondrial overexpression and knock-down of this complex without interfering with its nuclear or cytoplasmic functions were essential. By tagging CyclinB1/Cdk1 with MTS, we were able to achieve mitochondrial overexpression of this complex to study its mitochondrial targets as well as functions. Via tagging dominant-negative Cdk1 with MTS, inhibition of Cdk1 activity was accomplished particularly in the mitochondria. Potential mitochondrial targets of CyclinB1/Cdk1 complex were identified using a gel-based proteomics approach. Unlike traditional 2D gel analysis, we employed 2-dimensional difference gel electrophoresis (2D-DIGE) technology followed by phosphoprotein staining to fluorescently label differentially phosphorylated proteins in mitochondrial Cdk1 expressing cells. Identification of phosphoprotein spots that were altered in wild type versus dominant negative Cdk1 bearing mitochondria revealed the identity of mitochondrial targets of Cdk1. Finally, to determine the effect of CyclinB1/Cdk1 mitochondrial localization in cell cycle progression, a cell proliferation assay using a synthetic thymidine analogue EdU (5-ethynyl-2&#8242;-deoxyuridine) was used to monitor the cells as they go through the cell cycle and replicate their DNA. Altogether, we demonstrated a variety of approaches available to study mitochondrial localization and activity of a cell cycle kinase. These are advanced, yet easy to follow methods that will be beneficial to many cell biology researchers.

Here, we outline how to study mitochondrial localization of a (cell cycle) kinase, and how to determine its sub-mitochondrial location as well as potential mitochondrial substrates/targets. Forced expression of proteins into the mitochondria provides a useful tool for studying the functional consequences of mitochondrial localization of a protein of interest.

Although mitochondria possess their own transcriptional machinery, merely 1% of mitochondrial proteins are synthesized inside the organelle. The ...

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 Graphical User Interface for Software-assisted Tracking of Protein Concentration in Dynamic Cellular Protrusions

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex signaling mechanisms underlying filopodial initiation, elongation and subsequent stabilization or retraction, it is crucial to determine the spatio-temporal protein activity in these dynamic structures. To analyze protein function in filopodia, we recently developed a semi-automated tracking algorithm that adapts to filopodial shape-changes, thus allowing parallel analysis of protrusion dynamics and relative protein concentration along the whole filopodial length. Here, we present a detailed step-by-step protocol for optimized cell handling, image acquisition and software analysis. We further provide instructions for the use of optional features during image analysis and data representation, as well as troubleshooting guidelines for all critical steps along the way. Finally, we also include a comparison of the described image analysis software with other programs available for filopodia quantification. Together, the presented protocol provides a framework for accurate analysis of protein dynamics in filopodial protrusions using image analysis software.

We present a software solution for semi-automated tracking of relative protein concentration along the length of dynamic cellular protrusions.

Filopodia are dynamic, finger-like cellular protrusions associated with migration and cell-cell communication. In order to better understand the complex ...

Primary Culture of Rat Adrenocortical Cells and Assays of Steroidogenic Functions

The hormone which the adrenal cortex secretes is vital for animals against stress and diseases. The method described here is the procedure of primary cultured rat adrenal cells and related functional assays (immunofluorescence staining of lipid droplet surface protein, as well as corticosterone analysis). Unlike an in vivo model, the variation of interexperiments in adrenal monolayer cultures is less and the experimental condition is easy to control. Besides, the source of rats is also more stable than other animals, like bovine ones. There are also several human adrenal cell lines (NCI-H295, NCI-H295R, SW13, etc.) that can be used in adrenal studies. However, the steroid production of these lines will still be influenced by numerous factors, which include serum lot number, passage number, mutant/loss of distinct genes, etc. Except for lacking 17&#945;-hydroxylase, the primary culture of rat adrenocortical cells is a better and more convenient technique for studying adrenal physiology. In summary, primary rat adrenal cultures could be a good in vitro platform for researchers to investigate the mechanisms of the reagent of interest in the adrenal gland system.

The hormone secreted by the adrenal cortex is vital for animals against stress and diseases. Here, we present a protocol to culture the primary rat adrenal cells. It could be a good in vitro platform for investigating the mechanisms of the reagent of interest in adrenal steroidogenesis and lipid biosynthesis.

The hormone which the adrenal cortex secretes is vital for animals against stress and diseases. The method described here is the procedure of primary ...

Photostimulation by Femtosecond Laser Activates Extracellular-signal-regulated Kinase (ERK) Signaling or Mitochondrial Events in Target Cells

Direct control of cellular defined molecular events is important to life science. Recently, studies have demonstrated that femtosecond laser stimulation can simultaneously activate multiple cellular molecular signaling pathways. In this protocol, we show that through coupling femtosecond laser into a confocal microscope, cells can be stimulated precisely by the tightly-focused laser. Some molecular processes that can be simultaneously observed are subsequently activated. We present detailed protocols of the photostimulation to activate extracellular signal regulated kinase (ERK) signaling pathway in Hela cells. Mitochondrial flashes of reactive oxygen species (ROS) and other mitochondrial events can be also stimulated if focusing the femtosecond laser pulse on a certain mitochondrial tubular structure. This protocol includes pretreating cells before photostimulation, delivering the photostimulation by a femtosecond laser flash onto the target, and observing/identifying molecular changes afterwards. This protocol represents an all-optical tool for related biological researches.

Photostimulation by Femtosecond Laser Activates Extracellular-signal-regulated Kinase ERK Signaling or Mitochondrial Events in Target Cells

Tightly-focused femtosecond laser can deliver precise stimulation to cells by being coupled into a confocal microscopy enabling the real-time observation and photostimulation. The photostimulation can activate cell molecular events including ERK signaling pathway and mitochondrial flashes of reactive oxygen species.

Direct control of cellular defined molecular events is important to life science. Recently, studies have demonstrated that femtosecond laser stimulation ...