This work was supported by the National Institute of Aging (AG04418), National Institute on Drug Abuse, National Institute of Health-Karolinska Institutet Graduate Partnerships Program, Karolinska Institutet, Swedish Research Council, Swedish Brain Power, and Swedish Brain Foundation. Many thanks to Mattias Karlen and Dr. Giuseppe Coppotelli for creative support with Figure 1 and 2, respectively; Karin Pernold for technical assistance; and Drs. Barry J. Hoffer, Lars Olson, and Nils-G&ouml;ran Larsson for much helpful advice and discussion.

1. Tissue preparation for cryosectioning
<ol>
	<li>Sacrifice the animal by either cervical dislocation or decapitation, in accordance with available ethical permit.</li>
	<li>Quickly collect tissues of interest (e.g. brain), and rapidly freeze on dry ice (tissues may require freezing in isopentane or propane chilled with liquid nitrogen to obtain optimal morphology). Store tissues in aluminum foil at -80 &deg;C until ready to section.</li>
	<li>Embed frozen tissue in preparation for cryosectioning.</li>
	<li>Collect 14-&mu;m cryostat sections at -21 &deg;C (may need to adjust temperature &plusmn; 1-2 &deg;C). Thaw sections onto slides using heating block, and store slides without coverslipping at -20 &deg;C until ready to use.</li>
</ol>
2. COX histochemistry
<ol>
	<li>Allow slides to dry at room temperature for 1 hour. Put slides in a slide-staining chamber with wet filter paper, cut into strips. To obtain consistent results in each experiment, it is recommended to process a maximum of ten slides per experiment to minimize time delays.</li>
	<li>Prepare under a chemical hood 1X DAB, 100 &mu;M cytochrome c in 0.1 M PBS pH=7.0. Vortex quickly.</li>
	<li>Add 2 &mu;g bovine catalase (2 &mu;g ml-1 or approximately 4 IU ml-1). Mix well by vortexing to break up all the grains of catalase.</li>
	<li>Apply 150- 200 &mu;L of incubation medium to each slide, use pipette tip to spread evenly onto all sections.</li>
	<li>Incubate the slides for 40 minutes at 37 &deg;C.</li>
	<li>Remove excess solution from the slides. Wash slides 4 times, 10 minutes each time, in 0.1 M PBS pH=7.0.</li>
	<li>Return slides to slide-staining chamber with wet paper strips.</li>
</ol>
3. SDH histochemistry
<ol>
	<li>Prepare under a chemical hood 1.5 mM NBT, 130 mM sodium succinate, 0.2 mM PMS, and 1.0 mM sodium azide in 0.1 M PBS pH=7.0. Take caution to shield the PMS from light. Vortex quickly.</li>
	<li>Apply 150-200 &mu;L of incubation medium to each slide, use pipette tip to spread evenly onto all sections.</li>
	<li>Incubate the slides for 40 minutes at 37 &deg;C.</li>
	<li>Remove excess solution from the slides. Wash slides 4 times, 10 minutes each time, in 0.1 M PBS pH=7.0.</li>
	<li>Dehydrate the slides for 2 minutes in the following concentrations of ethanol: 70%, 70%, 95%, 95%, 99.5%. Then allow 10 minutes in an additional 99.5% step.</li>
	<li>Place slides in xylene for 10 minutes. Mount with Entellan and coverslip. Allow the slides to dry overnight, or at least 1-2 hours in a ventilated area.</li>
</ol>
4. Determination of mitochondrial dysfunction
<ol>
	<li>The amount of mitochondrial dysfunction is indicated by the amount of cellular blue staining. To semi-quantify these amounts, slides should be coded and visualized under bright-field microscopy. Semi-quantification should be performed on a blind basis using a scale, for example from 0-4 (0, no blue staining; 4, only blue staining). It is best to perform this kind of semi-quantification on several sections from a given subject/animal to calculate a mean value for each subject/animal.</li>
	<li>Statistics should be performed using a non-parametric test, such as Mann-Whitney or Kruskal-Wallis.</li>
</ol>
5. Appropriate specificity controls
<ol>
	<li>For specificity controls for COX activity, repeat "COX histochemistry" steps, and add 2.5 mM sodium azide, a terminal respiratory chain inhibitor.</li>
	<li>For specificity controls for SDH activity, repeat "SDH histochemistry" steps with the removal of sodium succinate and the addition of 50 mM malonate, a competitive inhibitor of SDH.</li>
	<li>Wash and dehydrate sections in an ethanol series, and then mount and coverslip the slides as described in steps 3.4 - 3.6.</li>
</ol>
6. Representative Results:
The overall scheme of the COX/SDH double-labeling histochemical assay is illustrated in Figure 2. Representative examples of appropriate COX/SDH double-labeling histochemistry in brain sections from wild-type and prematurely aging mtDNA mutator mice are shown in Figure 3. The dark brown staining in wild-type mice (Figure 3, left panel) showed normal COX activity. Cells with respiratory chain deficiencies, indicated by the blue staining, were revealed in 12 week-old mtDNA mutator mice, and these deficiencies became more widespread as mtDNA mutator mice aged to 46 weeks (Figure 3, center and right panel).
Examples of inappropriate COX/SDH double-labeling in brain sections from wild-type mice due to insufficient COX labeling are shown in Figure 4. Inadequate incubation time for the demonstration of COX activity, or reducing the availability of molecular oxygen by coverslipping the slide during incubation, resulted in a reduced deposition of the DAB reaction product, and thus allowed for formation of the blue formazan end product during the SDH incubation.
COX and SDH activities can also be investigated separately (Figure 5, left and center); however, the sequential labeling is helpful in identifying cells with COX deficiencies, due to the formation of the blue precipitate during the SDH incubation (Figure 3, center and right). Specificity controls for COX and SDH activities can also be done (Figure 5, right).
<img alt="Figure 1" src="/files/ftp_upload/3266/3266fig1.jpg" /> 
Figure 1. Mitochondrial respiratory Complexes I-V. The mitochondrial respiratory chain is located within the inner membrane and includes five complexes. The purpose of the respiratory chain is to transport electrons from Complex I to IV and in doing so it creates a proton gradient across the inner membrane used by Complex V (ATPase) to produce ATP. Red hexagons represent subunits encoded by mtDNA. White hexagons represent subunits encoded by nuclear DNA (note that Complex II is completely encoded from the nuclear genome). Thus, mutations in the mitochondrial genome could cause dysfunction of the respiratory chain due to mutations in the subunits of the respiratory chain complexes.
<img alt="Figure 2" src="/files/ftp_upload/3266/3266fig2.jpg" /> 
Figure 2. Flow chart of the COX/SDH double-labeling histochemical assay. Dissect the organs of interest, rapidly freeze the tissues on dry ice, and store them at - 80 &deg;C. Collect cryostat sections and keep at - 20 &deg;C until use. Allow sections to air-dry at room temperature for 1 hour. Prepare the incubation medium for COX histochemistry, apply it to the slides, and incubate for 40 minutes at 37 &deg;C. Wash the sections in PBS 4 times for 10 minutes each wash. Prepare the incubation medium for SDH histochemistry, apply it to the slides, and again incubate for 40 minutes at 37 &deg;C. Wash the sections again in PBS, dehydrate in an ethanol series, and then mount and coverslip the slides. The COX/SDH double-labeled sections are ready to view under bright-field microscopy within 1-2 hours.
<img alt="Figure 3" src="/files/ftp_upload/3266/3266fig3.jpg" /> 
Figure 3. Representative examples of COX/SDH double-labeling. Brain sections from wild-type and prematurely aging mtDNA mutator mice were sequentially labeled for COX and SDH activities. (Scale bar: 200 &mu;m.) Normal COX activity (indicated by dark brown color) was shown in hippocampus from wild-type mice (left). COX deficiencies (indicated by blue color) were revealed in hippocampus from mtDNA mutator mice (center and right). There was a further decrease in COX activity by 46 weeks of age in mtDNA mutator mice, suggesting widespread exacerbation of respiratory chain dysfunction. The observed mitochondrial dysfunction in mtDNA mutator mice 12 is caused by high levels of mtDNA point mutations as well as increased levels of linear deletions 5.
<img alt="Figure 4" src="/files/ftp_upload/3266/3266fig4.jpg" /> 
Figure 4. Examples of inappropriate COX/SDH double-labeling. Brain sections from wild-type mice were sequentially labeled for COX and SDH activities. (Scale bar: 200 &mu;m.) Inadequate incubation times (10 and 25 minutes) for the demonstration of COX activity resulted in a reduced deposition of the brown DAB reaction product, compared to the 40-minute incubation time (left and center). The shortened incubation times allowed for the formation of the blue formazan end product during the SDH incubation, misleadingly suggesting the presence of cells with COX deficiencies. Coverslipping the slides during the COX incubation also resulted in inaccurate formation and deposition of the DAB reaction product (right).
<img alt="Figure 5" src="/files/ftp_upload/3266/3266fig5.jpg" /> 
Figure 5. Individual COX and SDH labeling and specificity control. Brain sections from wild-type mice were separately labeled for COX and SDH activities, indicated by the dark brown color and the blue color, respectively (left and center). Although COX and SDH activities can be individually labeled, the sequential labeling has proved to be advantageous in locating cells with mitochondrial dysfunction. An example of a specificity control for COX and SDH activities in brain from a wild-type mouse showed absence of labeling (right). (Scale bar: 200 &mu;m.)

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No conflicts of interest declared.

The combined COX/SDH histochemical method enables the visualization of cells with mitochondrial dysfunction. This technique, with early studies dating back to 1968, remains popular, with many considering it the "gold standard" for identifying mitochondrial diseases in patients14,19,26,27. It is now frequently used to investigate mtDNA mutation-driven aging and aging-related disorders 12,13,18,20,21,24. The COX/SDH double-labeling method is often used in parallel with other techniques to identify spe...

<table border="1">
<tr>
<td>Name of the reagent</td>
<td>Company</td>
<td>Catalogue number</td>
<td>Comments (optional)</td>
</tr>
<tr>
<td>Dry Ice</td>
<td>AGA Gas AB</td>
<td>block form</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Isopentane (2-methylbutane)</td>
<td>Sigma-Aldrich</td>
<td>277258 
 CAS: 78-78-4</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Cyrostat embedding solution</td>
<td>Sakura Finetek</td>
<td>Tissue Tek 4583</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Cryostat</td>
<td>Microm</td>
<td>Microm Model HM 500M</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Slides</td>
<td>Thermo Scientific</td>
<td>Super Frost Plus 
 Menzel Gl&auml;ser 
 J1800AMWZ</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Cover glasses 
 Borosilicate glass</td>
<td>VWR International</td>
<td>16004-098</td>
<td>24 x 50 mm</td>
</tr>
<tr>
<td>Filter Paper</td>
<td>Munktell Filter AB</td>
<td>Quality: 1350 
 Article Number: 242 001</td>
<td>430 x 430 mm</td>
</tr>
<tr>
<td>3,3&prime;-diaminobenzidine tetrahydrochloride (DAB)</td>
<td>Sigma-Aldrich</td>
<td>Sigma Liquid Substrate System, D7304</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Cytochrome c (Type III, from equine heart)</td>
<td>Sigma-Aldrich</td>
<td>C2506 
 CAS: 9007-43-6</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Bovine catalase (from liver)</td>
<td>Sigma-Aldrich</td>
<td>C9322 
 CAS: 9001-05-2</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Nitroblue tetrazolium (NBT)</td>
<td>Sigma-Aldrich</td>
<td>N6876 
 CAS: 298-83-9</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Sodium succinate</td>
<td>Sigma-Aldrich</td>
<td>S2378 
 CAS: 6106-21-4</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Phenazine methosulfate (PMS)</td>
<td>Sigma-Aldrich</td>
<td>P9625 
 CAS: 299-11-6</td>
<td>PMS is light sensitive. Shield from light.</td>
</tr>
<tr>
<td>Sodium azide</td>
<td>Sigma-Aldrich</td>
<td>S8032 
 CAS: 26628-22-8</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Xylene</td>
<td>VWR International</td>
<td>EM-XX0060-4 </td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Entellan</td>
<td>VWR International</td>
<td>100503-870</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Malonate 
 (Malonic acid)</td>
<td>Sigma-Aldrich</td>
<td>M1296 
 CAS: 141-82-2</td>
<td>&nbsp;</td>
</tr>
</table>

visualization of mitochondrial respiratory function using cytochrome c oxidase / succinate dehydrogenase (cox/sdh) double-labeling histochemistry

Mitochondrial DNA (mtDNA) defects are an important cause of disease and may underlie aging and aging-related alterations 1,2. The mitochondrial theory of aging suggests a role for mtDNA mutations, which can alter bioenergetics homeostasis and cellular function, in the aging process 3. A wealth of evidence has been compiled in support of this theory 1,4, an example being the mtDNA mutator mouse 5; however, the precise role of mtDNA damage in aging is not entirely understood 6,7.
Observing the activity of respiratory enzymes is a straightforward approach for investigating mitochondrial dysfunction. Complex IV, or cytochrome c oxidase (COX), is essential for mitochondrial function. The catalytic subunits of COX are encoded by mtDNA and are essential for assembly of the complex (Figure 1). Thus, proper synthesis and function are largely based on mtDNA integrity 2. Although other respiratory complexes could be investigated, Complexes IV and II are the most amenable to histochemical examination 8,9. Complex II, or succinate dehydrogenase (SDH), is entirely encoded by nuclear DNA (Figure 1), and its activity is typically not affected by impaired mtDNA, although an increase might indicate mitochondrial biogenesis 10-12. The impaired mtDNA observed in mitochondrial diseases, aging, and age-related diseases often leads to the presence of cells with low or absent COX activity 2,12-14. Although COX and SDH activities can be investigated individually, the sequential double-labeling method 15,16 has proved to be advantageous in locating cells with mitochondrial dysfunction 12,17-21.
Many of the optimal constitutions of the assay have been determined, such as substrate concentration, electron acceptors/donors, intermediate electron carriers, influence of pH, and reaction time 9,22,23. 3,3'-diaminobenzidine (DAB) is an effective and reliable electron donor 22. In cells with functioning COX, the brown indamine polymer product will localize in mitochondrial cristae and saturate cells 22. Those cells with dysfunctional COX will therefore not be saturated by the DAB product, allowing for the visualization of SDH activity by reduction of nitroblue tetrazolium (NBT), an electron acceptor, to a blue formazan end product 9,24. Cytochrome c and sodium succinate substrates are added to normalize endogenous levels between control and diseased/mutant tissues 9. Catalase is added as a precaution to avoid possible contaminating reactions from peroxidase activity 9,22. Phenazine methosulfate (PMS), an intermediate electron carrier, is used in conjunction with sodium azide, a respiratory chain inhibitor, to increase the formation of the final reaction products 9,25. Despite this information, some critical details affecting the result of this seemly straightforward assay, in addition to specificity controls and advances in the technique, have not yet been presented.

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Visualization of Mitochondrial Respiratory Function using Cytochrome C Oxidase / Succinate Dehydrogenase COX/SDH Double-labeling Histochemistry

The cytochrome c oxidase/sodium dehydrogenase (COX/SDH) double-labeling method allows for direct visualization of mitochondrial respiratory enzyme deficiencies in fresh-frozen tissue sections. This is a straightforward histochemical technique and is useful in investigating mitochondrial diseases, aging, and aging-related disorders.

Visualization of Mitochondrial Respiratory Function using Cytochrome C Oxidase / Succinate Dehydrogenase (COX/SDH) Double-labeling Histochemistry

Mitochondrial DNA (mtDNA) defects are an important cause of disease and may underlie aging and aging-related alterations 1,2. The mitochondrial theory of ...

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Biology

36.5K Views.  Karolinska Institutet. Il citocromo c ossidasi / sodio deidrogenasi (COX / SDH) doppio metodo di etichettatura permette di visualizzare direttamente mitocondriale deficit enzimatici respiratori nei fresco congelato sezioni di tessuto. Questa è una tecnica semplice istochimiche ed è utile nelle indagini malattie mitocondriali, invecchiamento e patologie legate all'invecchiamento.

Video: Visualization of Mitochondrial Respiratory Function using Cytochrome C Oxidase / Succinate Dehydrogenase COX/SDH Double-labeling Histochemistry

Generation of Stable Transgenic C. elegans Using Microinjection

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1. The plasmid DNA rearranges to form extrachromosomal concatamers that are stably inherited, though not with the same efficiency as actual chromosomes 2. A gene of interest is co-injected with an obvious phenotypic marker, such as rol-6 or GFP, to allow selection of transgenic animals under a dissecting microscope. The exogenous gene may be expressed from its native promoter for cellular localization studies. Alternatively, the transgene can be driven by a different tissue-specific promoter to assess the role of the gene product in that particular cell or tissue. This technique efficiently drives gene expression in all tissues of C. elegans except for the germline or early embryo 3. Creation of transgenic animals is widely utilized for a range of experimental paradigms. This video demonstrates the microinjection procedure to generate transgenic worms. Furthermore, selection and maintenance of stable transgenic C. elegans lines is described.

This video demonstrates the technique of microinjection into the gonad of C. elegans to create transgenic animals.

Transgenic Caenorhabditis elegans can be readily created via microinjection of a DNA plasmid solution into the gonad 1.  The plasmid DNA rearranges to ...

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 ...

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

The isolated microvessel preparation is an ex vivo preparation that allows for examination of the different contributions of factors that control vessel diameter, and thus, perfusion resistance1-5. This is a classic experimental preparation that was, in large measure, initially described by Uchida et al.15 several decades ago. This initial description provided the basis for the techniques that was extensively modified and enhanced, primarily in the laboratory of Dr. Brian Duling at the University of Virginia6-8, and we present a current approach in the following pages. This preparation will specifically refer to the gracilis arteriole in a rat as the microvessel of choice, but the basic preparation can readily be applied to vessels isolated from nearly any other tissue or organ across species9-13. Mechanical (i.e., dimensional) changes in the isolated microvessels can easily be evaluated in response to a broad array of physiological (e.g., hypoxia, intravascular pressure, or shear) or pharmacological challenges, and can provide insight into mechanistic elements comprising integrated responses in an intact, although ex vivo, tissue. The significance of this method is that it allows for facile manipulation of the influences on the integrated regulation of microvessel diameter, while also allowing for the control of many of the contributions from other sources, including intravascular pressure (myogenic), autonomic innervation, hemodynamic (e.g., shear stress), endothelial dependent or independent stimuli, hormonal, and parenchymal influences, to provide a partial list. Under appropriate experimental conditions and with appropriate goals, this can serve as an advantage over in vivo or in situ tissue/organ preparations, which do not readily allow for the facile control of broader systemic variables. 
The major limitation of this preparation is essentially the consequence of its strengths. By definition, the behavior of these vessels is being studied under conditions where many of the most significant contributors to the regulation of vascular resistance have been removed, including neural, humoral, metabolic, etc. As such, the investigator is cautioned to avoid over-interpretation and extrapolation of the data that are collected utilizing this preparation. The other significant area of concern with regard to this preparation is that it can be very easy to damage cellular components such as the endothelial lining or the vascular smooth muscle, such that variable source of error can be introduced. It is strongly recommended that the individual investigator utilize appropriate measurements to ensure the quality of the preparation, both at the initiation of the experiment and periodically throughout the course of a protocol.

The ex vivo Isolated Skeletal Microvessel Preparation for Investigation of Vascular Reactivity

An ex vivo preparation is described for isolation of the largest gracilis muscle resistance arterioles for interrogation of both vascular responses to vasoactive stimuli and the assessment of basic structural properties via passive wall mechanics.

The isolated  microvessel preparation is an ex vivo preparation that allows for examination of the different contributions of  factors that control vessel ...

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.

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 ...

Visualization of Craniofacial Development in the sox10: kaede Transgenic Zebrafish Line Using Time-lapse Confocal Microscopy

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, convergence and extension of facial prominences, and maturation of the craniofacial skeleton. To study the contribution of the cranial neural crest to specific regions of the zebrafish palate a sox10: kaede transgenic zebrafish line was generated. Sox10 provides lineage restriction of the kaede reporter protein to the neural crest, thereby making the cell labeling a more precise process than traditional dye or reporter mRNA injection. Kaede is a photo-convertible protein that turns from green to red after photo activation and makes it possible to follow cells precisely. The sox10: kaede transgenic line was used to perform lineage analysis to delineate CNC cell populations that give rise to maxillary versus mandibular elements and illustrate homology of facial prominences to amniotes. This protocol describes the steps to generate a live time-lapse video of a sox10: kaede zebrafish embryo. Development of the ethmoid plate will serve as a practical example. This protocol can be applied to making a time-lapse confocal recording of any kaede or similar photoconvertible reporter protein in transgenic zebrafish. Furthermore, it can be used to capture not only normal, but also abnormal development of craniofacial structures in the zebrafish mutants.

Visualization of experimental data has become a key element in presenting results to the scientific community. Generation of live time-lapse recording of growing embryos contributes to better presentation and understanding of complex developmental processes. This protocol is a step-by-step guide to cell labeling via photoconversion of kaede protein in zebrafish.

Vertebrate palatogenesis is a highly choreographed and complex developmental process, which involves migration of cranial neural crest (CNC) cells, ...

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

RNA interference by feeding worms bacteria expressing dsRNAs has been a useful tool to assess gene function in C. elegans. While this strategy works well when a small number of genes are targeted for knockdown, large scale feeding screens show variable knockdown efficiencies, which limits their utility. We have deconstructed previously published RNAi knockdown protocols and found that the primary source of the reduced knockdown can be attributed to the loss of dsRNA-encoding plasmids from the bacteria fed to the animals. Based on these observations, we have developed a dsRNA feeding protocol that greatly reduces or eliminates plasmid loss to achieve efficient, high throughput knockdown. We demonstrate that this protocol will produce robust, reproducible knock down of C. elegans genes in multiple tissue types, including neurons, and will permit efficient knockdown in large scale screens. This protocol uses a commercially available dsRNA feeding library and describes all steps needed to duplicate the library and perform dsRNA screens. The protocol does not require the use of any sophisticated equipment, and can therefore be performed by any C. elegans lab.

Large-scale Gene Knockdown in C. elegans Using dsRNA Feeding Libraries to Generate Robust Loss-of-function Phenotypes

While dsRNA feeding in C. elegans is a powerful tool to assess gene function, current protocols for large scale feeding screens result in variable knockdown efficiencies. We describe an improved protocol for performing large scale RNAi feeding screens that results in highly efficacious and reproducible knockdown of gene expression.

RNA interference by feeding worms bacteria expressing dsRNAs has been a useful tool to assess gene function in C. elegans. While this strategy works well ...

Measuring Respiratory Function in Mice Using Unrestrained Whole-body Plethysmography

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative assessment of lung function in rodent models is an important tool in the development of future therapies. Commonly used techniques for assessing respiratory function including invasive plethysmography and forced oscillation. While these techniques provide valuable information, data collection can be fraught with artefacts and experimental variability due to the need for anesthesia and/or invasive instrumentation of the animal. In contrast, unrestrained whole-body plethysmography (UWBP) offers a precise, non-invasive, quantitative way by which to analyze respiratory parameters. This technique avoids the use of anesthesia and restraints, which is common to traditional plethysmography techniques. This video will demonstrate the UWBP procedure including the equipment set up, calibration and lung function recording. It will explain how to analyze the collected data, as well as identify experimental outliers and artefacts that results from animal movement. The respiratory parameters obtained using this technique include tidal volume, minute volume, inspiratory duty cycle, inspiratory flow rate and the ratio of inspiration time to expiration time. UWBP does not rely on specialized skills and is inexpensive to perform. A key feature of UWBP, and most appealing to potential users, is the ability to perform repeated measures of lung function on the same animal.

The assessment of respiratory physiology has traditionally relied upon techniques, which require restraint or sedation of the animal. Unrestrained whole-body plethysmography, however, provides precise, non-invasive, quantitative analysis of respiratory physiology in animal models. In addition, the technique allows repeated respiratory assessment of mice allowing for longitudinal studies.

Respiratory dysfunction is one of the leading causes of morbidity and mortality in the world and the rates of mortality continue to rise. Quantitative ...

High-resolution Respirometry to Assess Mitochondrial Function in Permeabilized and Intact Cells

A high-resolution oxygraph is a device for measuring cellular oxygen consumption in a closed-chamber system with very high resolution and sensitivity in biological samples (intact and permeabilized cells, tissues or isolated mitochondria). The high-resolution oxygraph device is equipped with two chambers and uses polarographic oxygen sensors to measure oxygen concentration and calculate oxygen consumption within each chamber. Oxygen consumption rates are calculated using software and expressed as picomoles per second per number of cells. Each high-resolution oxygraph chamber contains a stopper with injection ports, which makes it ideal for substrate-uncoupler-inhibitor titrations or detergent titration protocols for determining effective and optimum concentrations for plasma membrane permeabilization. The technique can be applied to measure respiration in a wide range of cell types and also provides information on mitochondrial quality and integrity, and maximal mitochondrial respiratory electron transport system capacity.

High-resolution respirometry is used to determine mitochondrial oxygen consumption. This is a straightforward technique to determine mitochondrial respiratory chain complexes&#39; (I-IV) respiratory rates, maximal mitochondrial electron transport system capacity, and mitochondrial outer membrane integrity.

A high-resolution oxygraph is a device for measuring cellular oxygen consumption in a closed-chamber system with very high resolution and sensitivity in ...

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Fluorescence microscopy is an indispensable tool for the direct assessment of mitochondrial structure and function in live cells and for studying the mitochondrial structure-function relationship, which is primarily modulated by the molecules governing fission and fusion events between mitochondria. This paper describes and demonstrates specific methods for studying mitochondrial structure and function in live as well as in fixed tissue in the model organism Drosophila melanogaster. The tissue of choice here is the Drosophila ovary, which can be isolated and made amenable for ex vivo live confocal microscopy. Furthermore, the paper describes how to genetically manipulate the mitochondrial fission protein, Drp1, in Drosophila ovaries to study the involvement of Drp1-driven mitochondrial fission in modulating the mitochondrial structure-function relationship. The broad use of such methods is demonstrated in already-published as well as in novel data. The described methods can be further extended towards understanding the direct impact of nutrients and/or growth factors on the mitochondrial properties ex vivo. Given that mitochondrial dysregulation underlies the etiology of various diseases, the described innovative methods developed in a genetically tractable model organism, Drosophila, are anticipated to contribute significantly to the understanding of the mechanistic details of the mitochondrial structure-function relationship and to the development of mitochondria-directed therapeutic strategies.

Studying Mitochondrial Structure and Function in Drosophila Ovaries

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial functionality. Specific methods for studying mitochondrial structure and function in live and fixed Drosophila ovaries are described and demonstrated in this paper.

Analysis of the mitochondrial structure-function relationship is required for a thorough understanding of the regulatory mechanisms of mitochondrial ...

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.