R.J.M. is supported by an AMC PhD Scholarship. This research was supported by the Dutch Cancer Society (KWF grant UVA 2014-6839). The authors thank Dr. A. Jonker for his help with the writing of the protocol.

This protocol follows the guidelines on the use of human material of the Medical-Ethical Review Board of the Academic Medical Center.
1. Preparation of Cells or Tissue Sections
<ol>
	<li>Cells
	<ol>
		<li>Trypsinize cells from culture dishes using 1 mL of 0.25% trypsin-EDTA for 5 min at 37&#160;&#176;C in a 10 mm Petri dish and bring cells in a 37&#160;&#176;C suspension of 50,000-200,000 cells/mL, depending on the size of the cells.</li>
		<li>Attach 200 &#181;L of the cell suspension onto microscopy slides using cytospin funnels in a cytocentrifuge at 20 x g for 5 min at room temperature.</li>
		<li>Air-dry the cytospins for 24 h at room temperature. This does not affect dehydrogenase activity.5</li>
	</ol>
	</li>
	<li>Tissue Sections
	<ol>
		<li>Extract tissue from patients or animals according to ethical permits. 10 mm3 of tissue is sufficient for multiple experiments.</li>
		<li>Put the pieces of tissue into small vented 2 mL plastic vials and snap-freeze the vials containing the piece of tissue in liquid nitrogen.</li>
		<li>Store the vials containing tissue pieces at &#8722;80 &#176;C until further processing.</li>
		<li>Use a gel-like medium called optimal cutting temperature (OCT) to mount the tissue sample on a chuck that freezes down to -20 &#176;C to -25 &#176;C rapidly, so that water cannot form crystals.</li>
		<li>Use a cryostat for cutting and first trim the block to the desired level (ideally half the width of a microscopy glass slide, e.g. 5 mm x 5 mm).</li>
		<li>Upon sectioning, use brushes and an anti-roll plate to prevent the sections from curling upwards.</li>
		<li>Cut tissue samples into 6-10 &#181;m-thick sections at a slow but constant speed (1 s per section). When a section is correctly cut, it remains flat on the microtome knife under the anti-roll plate.</li>
		<li>Pick up 1-3 sections on microscopical glass slides and store at &#8722;80 &#176;C until use. The number of sections prepared depends on the size of the tissue sample and the desired thickness of the sections.</li>
	</ol>
	</li>
</ol>
2. Enzyme Histo/Cytochemistry for Soluble Dehydrogenases
<ol>
	<li>Preparing Reaction Buffer.
	<ol>
		<li>Prepare 100 mL of phosphate buffer of the correct pH (see Table 1; every enzyme has an optimal pH at which its activity is the highest). For example, mix solutions of 0.1 M KH2PO4 (acidic) and 0.1 M NA2HPO4 (basic) until a buffer with the correct pH is made.</li>
		<li>Weigh 18 g of polyvinyl alcohol (PVA) into the phosphate buffer under a fume hood as PVA is dusty and toxic.
		<ol>
			<li>Dissolve the 18% w/v PVA in the phosphate buffer at 100 &#176;C by warming the buffer au bain marie (put the glass bottle in boiling water) and stirring until the PVA is completely dissolved. The solution will be transparent then, with small air bubbles that are caused by the stirring. It typically requires ~15 min for the PVA to dissolve. 
			NOTE: The 18% w/v PVA in phosphate buffer is viscous and can be transferred to 15 mL reaction buffer tubes using a wide pipette. Short-term storage of 18% w/v PVA in phosphate buffer should occur at &#8805;40 &#176;C and long-term storage (up to 2 weeks) at &#8805; 60 &#176;C. 250 &#181;L of 18% w/v PVA in phosphate buffer is typically required for a cytospin preparation, while 500 &#181;L is required for a tissue section.</li>
		</ol>
		</li>
		<li>Prepare tetrazolium salt (nitroBT) solution. For every 1 mL of incubation medium, 40 &#181;L of nitroBT solution is required, consisting of 5 mg of nitroBT (a yellow powder) dissolved in a mix of 20 &#181;L dimethylformamide (DMF) and 20 &#181;L of ethanol, which will be a light yellow transparent solution. 
		NOTE: Because the stability of nitroBT is the lowest of all necessary reagents, it is recommended to prepare the nitroBT stock solution last and dissolve the nitroBT into the reaction medium first.
		<ol>
			<li>Dissolve nitroBT in DMF and ethanol by heating it in a glass vial during a short period of time (~10 s) using a Bunsen burner. Shake the vial during heating and do not leave the vial too long in the flame to prevent evaporation.</li>
			<li>Make 10% extra nitroBT solution to allow for the evaporation of DMF and ethanol during the dissolving process.</li>
		</ol>
		</li>
		<li>Shield the electron carriers phenazine methosulfate (PMS) or (1-methoxy)-phenazine methosulfate (mPMS), and incubation media containing m(PMS) from direct light by wrapping tinfoil around the glass storage vial.</li>
		<li>Prepare reagent solutions according to Table 1 by dissolving them in distilled H2O. Some reagents perish rapidly, so prepare them as shortly before the experiment as possible. Keep reagent solutions on ice or at 4 &#176;C until use.</li>
		<li>Prepare incubation media as shown in Table 1 and homogenize the solution after each step by gentle stirring. Avoid the generation of air bubbles in the incubation medium by stirring continuously and keeping the spatula below the surface of the incubation medium while stirring.</li>
	</ol>
	</li>
	<li>Applying Incubation Medium to Tissue Sections
	<ol>
		<li>Pre-warm microscopy slides with cytospins or tissue sections attached at 37 &#176;C in an immunohistochemistry incubation chamber for 15 min. Submerge the bottom of the immunohistochemistry incubation chamber with warm water to maintain a constant temperature in the incubator.</li>
		<li>Carefully break or saw off Pasteur pipettes at the transition from the narrow to the wide part. Regular Pasteur pipettes are too narrow to aspirate the viscous incubation medium, so this step is needed to make Pasteur pipettes wide enough for the aspiration of the incubation medium.</li>
		<li>Apply 250-500 &#181;L of the appropriate incubation medium to each cell preparation or tissue section on microscopy slides using the wide part of a Pasteur pipette.</li>
		<li>Use the pipette tip to spread out the incubation medium evenly. Disregard small air bubbles in the incubation medium, because these will float to the surface and will not interfere with the enzymatic reaction in the cell preparation or tissue section on the microscopy slide.</li>
		<li>Incubate the cell preparations or tissue sections on the microscopy slides in a 37 &#176;C incubator (e.g. a tissue culture incubator) for a pre-specified duration, depending on the enzyme kinetics and its expression in a particular cell preparation or tissue (Table 1). Keep the lid of the immunohistochemistry incubation chamber closed to maintain a humid environment to prevent the reaction buffer from drying out.</li>
	</ol>
	</li>
	<li>Washing the Microscopy Slides
	<ol>
		<li>Tap off excess incubation medium on a tissue.</li>
		<li>Mechanically rinse off the incubation medium from the microscopy slides in 60 &#176;C phosphate buffer, pH 5.3, by moving the microscopy slides up and down in the buffer. This stops the enzymatic reaction.</li>
		<li>Wash the microscopy slides by keeping them in 60 &#176;C phosphate buffer, pH 5.3 for 20 min.</li>
		<li>Mechanically wash the microscopy slides in 60 &#176;C tap water.</li>
		<li>Wash the microscopy slides by keeping them in 60 &#176;C tap water for 20 min.</li>
		<li>Let the water and slides cool down by slowly mixing room temperature tap water with the 60 &#176;C tap water over the course of 1 minute.</li>
		<li>Perform a final mechanical washing step in room temperature distilled water.</li>
	</ol>
	</li>
	<li>Enclose the Stained Cytospins or Tissue Sections on Microscopy Slides
	<ol>
		<li>Pre-warm a slide warmer at 50-60 &#176;C.</li>
		<li>Carefully dry the back side of the microscopy slides using a paper towel and place them on the slide warmer to dry the top side of the microscopy slide.</li>
		<li>When the slides are dry, enclose cell preparations or tissue sections on microscopy slides using glycerol jelly mounting medium and coverslips.</li>
		<li>Save the cell preparations or tissue sections microscopy slides in the dark in a refrigerator at 4 &#176;C or immediately proceed with image acquisition.</li>
	</ol>
	</li>
</ol>
3. Image Acquisition
<ol>
	<li>Record images with a scientific monochrome CCD camera with sufficient dynamic range (at least 8-bit but preferably 10-bit or 12-bit), fixed gain and a linear response.</li>
	<li>Select a proper objective (20X-63X for cytospins and 10-63X for tissue sections).</li>
	<li>Reduce glare by narrowing the field diaphragm so that it is slightly larger than the field of view.</li>
	<li>Use monochromatic light by applying a&#160;10 nm wide bandpass inference filter near the isobestic wavelength of the formazan precipitate7 in combination with an infrared blocking filter (see Materials Table). 
	NOTE: The maximum isobestic absorbance of nitroBT is at 585 nm.</li>
	<li>Optimize illumination to ensure that the entire grey level range of the camera is used (i.e., set the illumination level so that maximum illumination is achieved without over-exposing when recording a blank microscope slide).</li>
	<li>Calibrate measured grey levels to known absorbance (or optical density [OD]) values. To this end, capture grey scale images of at least 10 different steps of a calibration slide or step tablet with known OD values (see step 4.1.1), either commercially available (see Materials Table) or measure a custom-made grey scale wedge step tablet by a densitophotometer and use the results to calibrate measured grey values to known absorbance values.</li>
	<li>Capture photomicrographs of cells or tissue sections of interest.</li>
</ol>
4. Image Analysis
<ol>
	<li>Before image analysis, calibrate the ImageJ software for absorbance measurements.
	<ol>
		<li>Measure the absorbance (OD) of photomicrographs of at least 10 calibration areas with known absorbance parameters in the calibration slide or step tablet with known OD values. In ImageJ, use the Analyze &#8594; Measure menu or Ctrl+M hotkey to measure a selected area.</li>
		<li>Start the absorbance calibration procedure by going to Analyze &#8594; Calibration. Choose function &#34;Rodbard (NIH Image)&#34;. In the left column of the window that opens, the results of grey level measurements from each step of the calibration slide/step tablet performed in 4.1.1 are already present. If not, copy them from the ImageJ results screen. In the right column, enter each corresponding known absorbance value as printed on the certificate you received with the calibrated step tablet. Tick boxes &#34;Global calibration&#34; and &#34;Show plot&#34; and press &#34;OK&#34;.</li>
		<li>For quality control, make sure the R2 of the Rodbard function is &#62;0.99. If it is &#60;0.99, check whether the calibration slides were photographed and measured correctly and whether the known absorbance parameters were entered correctly.</li>
		<li>ImageJ is now calibrated until you close the program (hence &#34;Global calibration&#34;). For absorbance measurements, a calibrated ImageJ session always converts the observed absorbance means to values between 0 and 1, where 0 and 1 are the asymptotes of zero and full absorbance, respectively.</li>
	</ol>
	</li>
	<li>For cytospins, select &#62;100 single cells in a random manner. For tissue sections, select one or more fields of interest of a fixed length and width in all sections.
	<ol>
		<li>Project a raster over the photomicrographs to assist in unbiased cell selections. In ImageJ, project a grid over an image via the Plugins &#8594; Analyze &#8594; Grid menu.</li>
		<li>In ImageJ, use the Elliptical Tool to select single cells, and use the Rectangle Tool to select tissue regions.</li>
	</ol>
	</li>
	<li>Measure the absorbance and area of the selected cells or tissue regions. The optical density and area are called &#34;Mean&#34; and &#34;Area&#34; in the ImageJ results spreadsheet, respectively. 
	NOTE: The total absorbance of a cell or tissue region (if you have selected several tissue regions with different sizes) is equal to the sum of all separated pixel absorbances. If a cell is imaged by 1000 pixels, then the total absorbance is given by the sum of 1000 absorbances and the mean absorbance is given by total absorbance/1000. This procedure also avoids distributional error.</li>
	<li>Determine the average total absorbance of &#62;30 cells or tissue regions from the control slide(s), which contains a sample that was exposed to control incubation medium in the absence of substrate but in the presence of cofactors. This control absorbance is used to control for non-specific enzyme activity staining and absorbance artefacts induced by the used microscopy slides, mounting medium, cover slip or microscope.</li>
	<li>Subtract the average control absorbance from all total absorbance measurements of samples that were exposed to incubation medium with the same concentration of cofactor (but different concentrations of substrate and/or inhibitor). This gives the corrected total absorbance of a cell or tissue region.</li>
	<li>Statistically compare the average corrected total absorbances using the Student&#39;s T-test or one-way ANOVA test, depending on your experimental design.</li>
	<li>Formazan absorbance can be converted into enzyme activity (&#181;mol converted substrate per mL per min) using the law of Lambert-Beer: c=A/(&#1108;&#215;d), where c is the concentration of formazan in the reaction medium, A is the measured absorbance in ImageJ after calibration; &#1108; is the extinction coefficient of formazan (16.000 at 585 nm)8 and d is the light traveling distance, which is equal to the average cell thickness or nominal section cutting thickness (6-10 &#181;m in this protocol). 
	NOTE: After drying, the tissue section thickness decreases ~50% from the nominal section cutting thickness, but this is not reflected in the calculation above because the nominal section cutting thickness is biologically the most relevant. The average cell thickness and sphere dimensions of cell preparates adhered to microscopy glass slides can be determined using confocal microscopy or wide-field microscopy, for which protocols can be found elsewhere.9,10</li>
</ol>

<ol>
	<li>Pavlova, N. N., Thompson, C. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Emerging+Hallmarks+of+Cancer+Metabolism.">The Emerging Hallmarks of Cancer Metabolism.</a> Cell Metab. 23 (1), 27-47 (2016).</li><li>Van Noorden, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Imaging+enzymes+at+work:+metabolic+mapping+by+enzyme+histochemistry.">Imaging enzymes at work: metabolic mapping by enzyme histochemistry.</a> J Histochem Cytochem. 58 (6), 481-497 (2010).</li><li>Van Noorden, C. J. F., Mcmanus, L. M., Mitchell, R. N. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Pathobiology of Human Disease. , 3760-3774 (2014).</li><li>Chieco, P., Jonker, A., De Boer, B. A., Ruijter, J. M., Van Noorden, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Image+cytometry:+protocols+for+2D+and+3D+quantification+in+microscopic+images.">Image cytometry: protocols for 2D and 3D quantification in microscopic images.</a> Prog Histochem Cytochem. 47 (4), 211-333 (2013).</li><li>Van Noorden, C. J. F., Frederiks, W. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Enzyme histochemistry : a laboratory manual of current methods. , (1992).</li><li>Van Noorden, C. J. F., Butcher, R. G., Stoward, P. J., Pearse, A. G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Histochemistry, theoretical and applied. Vol 3.: enzyme histochemistry. , 355-432 (1991).</li><li>Van Noorden, C. J., Tas, J., Vogels, I. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytophotometry+of+glucose-6-phosphate+dehydrogenase+activity+in+individual+cells.">Cytophotometry of glucose-6-phosphate dehydrogenase activity in individual cells.</a> Histochem J. 15 (6), 583-599 (1983).</li><li>Butcher, R. G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+measurement+in+tissue+sections+of+the+two+formazans+derived+from+nitroblue+tetrazolium+in+dehydrogenase+reactions.">The measurement in tissue sections of the two formazans derived from nitroblue tetrazolium in dehydrogenase reactions.</a> Histochem J. 10 (6), 739-744 (1978).</li><li>Matsumoto, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Cell biological applications of confocal microscopy. , (2002).</li><li>Errington, R. J., White, N. S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Measuring+dynamic+cell+volume+in+situ+by+confocal+microscopy.">Measuring dynamic cell volume in situ by confocal microscopy.</a> Methods Mol Biol. 122, 315-340 (1999).</li><li>Molenaar, R. J., Radivoyevitch, T., Maciejewski, J. P., van Noorden, C. J., Bleeker, F. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+driver+and+passenger+effects+of+isocitrate+dehydrogenase+1+and+2+mutations+in+oncogenesis+and+survival+prolongation.">The driver and passenger effects of isocitrate dehydrogenase 1 and 2 mutations in oncogenesis and survival prolongation.</a> Biochim Biophys Acta. 1846 (2), 326-341 (2014).</li><li>Molenaar, R. J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Radioprotection+of+IDH1-Mutated+Cancer+Cells+by+the+IDH1-Mutant+Inhibitor+AGI-5198.">Radioprotection of IDH1-Mutated Cancer Cells by the IDH1-Mutant Inhibitor AGI-5198.</a> Cancer Res. 75 (22), 4790-4802 (2015).</li><li>Khurshed, M., Molenaar, R. J., Lenting, K., Leenders, W. P., van Noorden, C. J. F. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=In+silico+gene+expression+analysis+reveals+glycolysis+and+acetate+anaplerosis+in+IDH1+wild-type+glioma+and+lactate+and+glutamate+anaplerosis+in+IDH1-mutated+glioma.">In silico gene expression analysis reveals glycolysis and acetate anaplerosis in IDH1 wild-type glioma and lactate and glutamate anaplerosis in IDH1-mutated glioma.</a> Oncotarget. 8 (30), 49165-49177 (2017).</li><li>Alberts, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Molecular biology of the cell. , (2015).</li><li>Berg, J. M., Tymoczko, J. L., Gatto, G. J., Stryer, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Biochemistry. , (2015).</li><li>Webb, E. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Enzyme nomenclature 1992 : recommendations of the Nomenclature Committee of the International Union of Biochemistry and Molecular Biology on the nomenclature and classification of enzymes. , (1992).</li><li>Botman, D., Tigchelaar, W., Van Noorden, C. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Determination+of+Phosphate-activated+Glutaminase+Activity+and+Its+Kinetics+in+Mouse+Tissues+using+Metabolic+Mapping+(Quantitative+Enzyme+Histochemistry).">Determination of Phosphate-activated Glutaminase Activity and Its Kinetics in Mouse Tissues using Metabolic Mapping (Quantitative Enzyme Histochemistry).</a> J Histochem Cytochem. 62 (11), 813-826 (2014).</li><li>van Lith, S. A., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glutamate+as+chemotactic+fuel+for+diffuse+glioma+cells:+are+they+glutamate+suckers?.">Glutamate as chemotactic fuel for diffuse glioma cells: are they glutamate suckers?.</a> Biochim Biophys Acta. 1846 (1), 66-74 (2014).</li></ol>

The authors have nothing to disclose.

Research on cellular metabolism is currently experiencing a renaissance because researchers realize now that metabolic effects are crucial for the pathogenesis and treatment of many diseases.1 Furthermore, metabolic research is aided by an increasing availability of techniques that provide this field of research with more tools than ever, including mass spectrometry, radioisotopic labeling and nuclear magnetic resonance spectrometry. Metabolic mapping is by no means a novel technique, but its capacity to give integrated information on the activity of enzymes in an almost true-to-nature situation makes this technique more relevant than ever.2
The cofactors NAD+ and NADP+ are found in all living cells, which indicates that dehydrogenases arose very early during evolution and have key roles in cellular metabolism.14,15 Currently, there are 523 enzyme-catalyzed reactions that are classified as dehydrogenases and occur across all species.16 Theoretically, the activity of all different dehydrogenase reactions can be separately investigated by tweaking the metabolic mapping protocol described here. Every enzymatic reaction is unique by means of the substrate and cofactors that are needed for its activity. Therefore, the activity of each enzymatic reaction can be determined using a metabolic mapping experiment with the adequate substrate and cofactors in the reaction medium. However, some enzyme isoforms differ in their subcellular localization, e.g. one isoform catalyzes a reaction in the cytoplasm whereas another isoform functions in the mitochondria. A notable example is IDH1 and IDH2, of which the former is cytoplasmic and the latter is mitochondrial and which are two different proteins encoded by two different genes.11 1-methoxy-5-methylphenazinium methylsulfate (methoxy-PMS) and 5-methylphenazinium methylsulfate (PMS) can both be used as electron carriers for these experiments. The former does not pass mitochondrial membranes, the latter does. Therefore, investigations on mitochondrial enzymes (e.g. IDH2) should use PMS whereas investigations on cytosolic enzymes (e.g. IDH1) should use mPMS.
As with many techniques, variations of this protocol, such as using different types of tetrazolium salts, excluding the addition of PMS and sodium azide, using an aqueous mounting medium, and varying incubation and rinsing times, do exist and may work equally well. The application of metabolic mapping with tetrazolium salts is not limited to dehydrogenases. With small modifications to the protocol, it can also be used for the assessment of the activity of enzymes that function directly upstream of a dehydrogenase in a metabolic pathway. We have previously described this principle for the glutaminase enzyme,17 which functions directly upstream of glutamate dehydrogenase in the glutaminolysis pathway.18 In theory, the same principle can be applied to other enzymes that function directly downstream or upstream of a dehydrogenase e.g. aconitase, which lies directly upstream of IDH2 and IDH3 in the tricarboxylic acid (TCA) cycle. Another simple variation of the concentrations described in Table 1 is by means of making incubation medium vials with various concentrations of substrate, cofactor and/or inhibitor. This allows the generation of dose-activity curves as a function of substrate, cofactor and/or inhibition concentration.
Technically speaking, metabolic mapping experiments do not facilitate unbiased observations of in situ enzyme activity, because researchers have to choose a substrate and cofactor concentration that may not reflect the substrate and cofactor levels that are present in situ. Furthermore, the use of viscous 18% PVA in the reaction medium serves to keep macromolecules intact, in place and in conformation but prohibits effective diffusion of low molecular-weight reagents through the reaction medium.3,5 Therefore, the determined enzyme activities in the experiments described here that use supraphysiological substrate concentrations do not reflect the in vivo situation at a given substrate concentration but are suitable for intra-experimental comparisons. Thus, the outcome of metabolic mapping experiments is the production capacity (maximum activity) of an enzymatic reaction in the context of the substrate and cofactor levels that are present in situ. Moreover, the use of various concentrations of substrate and/or cofactors and multiple samples allows unbiased comparison of the maximum production capacity (Vmax) and affinity (Km) of an enzyme for a substrate/cofactor in cell preparations, tissues or tissue regions. These parameters are the result of the sum of enzyme protein expression, post-translational modifications and the effect of a crowded microenvironment on the activity of enzymes. Therefore, metabolic mapping is still a better reflection of enzyme activity than experiments that determine protein expression or purified enzyme activity in vitro.

An essential cornerstone of cellular physiology is metabolism. Metabolism provides cells with the energy needed for all physiological processes, delivers the building blocks for macromolecular biosynthesis, and regulates cellular homeostasis with respect to waste products, toxic molecules and the disassembly and recycling of unnecessary or dysfunctional cellular components. Enzymes catalyze nearly all vital cellular chemical reactions and therefore are the motor units in the physiology of cells.1,2
The activity of enzymes is tightly regulated at many levels and therefore quantitative enzyme histochemistry and cytochemistry (also called metabolic mapping) is the preferred method to study enzyme activity in situ. At the transcriptional level, gene expression into mRNA is regulated. The effect of the regulation of transcription can be determined using quantitative mRNA assays, such as microarrays or direct RNA sequencing, or qualitative mRNA assays, such as in-situ hybridization which gives information on the (sub)cellular localization of mRNA molecules. This makes it possible to appreciate relative differences in transcriptional activity between cells. However, the interpretation and validity of these mRNA assays with respect to metabolic activity is complicated because regulation also occurs at the mRNA level, where sequence and stability of mRNA molecules are edited after it is transcribed from DNA. This editing regulates protein isoform translation and protein translation quantity at the ribosomes. In addition, the process of protein translation is controlled and ultimately affects enzyme expression and therefore activity. The combined effects of the aforementioned regulatory steps can be appreciated using quantitative protein expression assays, such as Western Blotting or reverse phase protein lysate microarrays, or qualitative protein expression assays, such as immunocytochemistry and immunohistochemistry, but these techniques fail to incorporate downstream regulatory effects such as post-translational modification of proteins and the functional regulation of enzymes in their crowded microenvironment. Moreover, the expression of an enzyme may poorly correlate with its activity, so that activity measurements of a purified enzyme in cell or tissue homogenates or in diluted solutions are widely used for enzyme activity studies. However, these experiments fail to replicate the influence of a crowded compartmentalized cytoplasm or organelle on the activity of an enzyme. Furthermore, all aforementioned techniques determine either the quantity or the localization of mRNA or enzyme expression, but are unable to give comprehensive information on both of these aspects of enzyme expression, let alone the integration of this information with enzyme activity determinations.2,3,4
Metabolic mapping allows the appreciation of all of the aforementioned variables that determine the activity of an enzyme. What is more, metabolic mapping is a form of living cell or tissue imaging with cells and tissues that are kept intact during analysis to generate an almost true-to-nature situation to generate data of activity of enzymes. It produces images that facilitate both the detailed appreciation of the location of enzyme activity as well as robust quantification of enzyme activity within a cell or tissue compartment.3,4 The protocols described here for metabolic mapping of activity of dehydrogenases are based on the laboratory manual of all available enzyme histochemical methods,5 on the principles of quantitative enzyme histochemistry,6 and on image analysis of quantitative metabolic mapping.4
Dehydrogenases are enzymes that perform redox reactions to reduce canonical cofactors such as NAD+, NADP+ and FAD to NADH, NADPH and FADH2, respectively. Intact cells or cryostat tissue sections that are not chemically fixed are incubated in a reaction medium which contains, among other reagents, the substrate and cofactors of a specific dehydrogenase and a tetrazolium salt. Subsequently, that dehydrogenase performs its catalytic activity and reduces, for example, NADP+ to NADPH. Via an electron carrier, 2 NADPH molecules ultimately reduce 1 water-soluble semi-colorless tetrazolium salt molecule into 1 water-insoluble blue formazan molecule that immediately precipitates at the site of the dehydrogenase. Therefore, the absorbance of precipitated formazan is a direct measure of the local activity of a dehydrogenase and can be observed using light microscopy or quantified using monochromatic light microscopy and image analysis. The quantitative aspect of this protocol enables researchers to draw statistical conclusions from these assays. In addition, this quantification facilitates the determination of in situ kinetic enzyme parameters, such as the maximal enzyme activity (Vmax) and affinity of a substrate for an enzyme (Km).3,4,5,6
When planning a metabolic mapping experiment, it is important to realize that per experiment, a maximum of fifty glass slides with adhering cell preparations or tissue sections are recommended to minimize time delays during incubation in order to obtain consistent results. It is possible to process more slides and/or medium compositions when these protocol steps are carried out cooperatively by more than one experimenter. Furthermore, some variability exists between experiments. Therefore, identical experiments should be repeated at least 3 times with cytospins from each cell preparation or sections from each tissue sample and appropriate controls should be included in each experiment. Always prepare a control incubation medium in the absence of substrate but in the presence of cofactors to control for non-specific enzyme activity staining. The most representative results are obtained in experiments where different substrate/cofactor concentrations are applied to tissue sections or cell preparations from the same sample, so that the experimenter can perform analyses of enzyme kinetics using dose-activity curves.

<table border="0" cellpadding="0" cellspacing="0" width="972">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Adenosine 5&#8242;-diphosphate sodium salt</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">A2754</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">D-Glucose 6-phosphate sodium salt</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">G7879</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:343px;">Disodium hydrogen phosphate (NA2HPO4)</td>
			<td style="width:343px;">Merck Millipore</td>
			<td align="right" style="width:119px;">106559</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">di-Sodium succinate</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">W327700</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">DL-Isocitric acid trisodium salt hydrate</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">I1252&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">D-Malic acid</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td align="right" style="width:119px;">2300</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Glycerol Gelatin</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">GG1</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">L-Glutamic acid</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">G1251</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Lithium 6-phospho-D-galactonate</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td align="right" style="width:119px;">55962</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Magnesium chloride</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">M8266</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">methoxy-Phenazine methosulfate</td>
			<td style="width:343px;">Serva</td>
			<td style="width:119px;">28966.01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Nicotinamide adenine dinucleotide</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">N0505&#160;</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Nicotinamide adenine dinucleotide phosphate</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">NADP-RO</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Nitrotetrazolium Blue chloride</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">N6876</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Phenazine methosulfate</td>
			<td style="width:343px;">Serva</td>
			<td style="width:119px;">32030.01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Polyvinyl alcohol, fully hydrolyzed</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">P1763</td>
			<td></td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:343px;">Potassium dihydrogen phosphate (KH2PO4)</td>
			<td style="width:343px;">Merck Millipore</td>
			<td align="right" style="width:119px;">104873</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Sodium azide</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">S2002</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Sodium L-lactate</td>
			<td style="width:343px;">Sigma-Aldrich</td>
			<td style="width:119px;">L7022</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Bandpass filter</td>
			<td style="width:343px;">Edmund optics</td>
			<td style="width:119px;"></td>
			<td>https://www.edmundoptics.de/optics/optical-filters/bandpass-filters/Hard-Coated-OD-4-10nm-Bandpass-Filters/</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:343px;">Grey-step wedge</td>
			<td style="width:343px;">Stouffer Industries</td>
			<td style="width:119px;"></td>
			<td>http://www.stouffer.net/TransPage.htm</td>
		</tr>
	</tbody>
</table>

metabolic mapping: quantitative enzyme cytochemistry and histochemistry to determine the activity of dehydrogenases in cells and tissues

Altered cellular metabolism is a hallmark of many diseases, including cancer, cardiovascular diseases and infection. The metabolic motor units of cells are enzymes and their activity is heavily regulated at many levels, including the transcriptional, mRNA stability, translational, post-translational and functional level. This complex regulation means that conventional quantitative or imaging assays, such as quantitative mRNA experiments, Western Blots and immunohistochemistry, yield incomplete information regarding the ultimate activity of enzymes, their function and/or their subcellular localization. Quantitative enzyme cytochemistry and histochemistry (i.e., metabolic mapping) show in-depth information on in situ enzymatic activity and its kinetics, function and subcellular localization in an almost true-to-nature situation.
We describe a protocol to detect the activity of dehydrogenases, which are enzymes that perform redox reactions to reduce cofactors such as NAD(P)+ and FAD. Cells and tissue sections are incubated in a medium that is specific for the enzymatic activity of one dehydrogenase. Subsequently, the dehydrogenase that is the subject of investigation performs its enzymatic activity in its subcellular site. In a chemical reaction with the reaction medium, this ultimately generates blue-colored formazan at the site of the dehydrogenase&#39;s activity. The formazan&#39;s absorbance is therefore a direct measure of the dehydrogenase&#39;s activity and can be quantified using monochromatic light microscopy and image analysis. The quantitative aspect of this protocol enables researchers to draw statistical conclusions from these assays.
Besides observational studies, this technique can be used for inhibition studies of specific enzymes. In this context, studies benefit from the true-to-nature advantages of metabolic mapping, giving in situ results that may be physiologically more relevant than in vitro enzyme inhibition studies. In all, metabolic mapping is an indispensable technique to study metabolism at the cellular or tissue level. The technique is easy to adopt, provides in-depth, comprehensive and integrated metabolic information and enables rapid quantitative analysis.

Several protocols to prepare incubation medium are shown to investigate the activity of a particular dehydrogenase. For each dehydrogenase, dissolve the listed reagents in the PVA solution while maintaining a temperature of at least 37 &#176;C. The reagents should be dissolved in the order listed in the table, i.e. nitroBT is always dissolved first and (m)PMS is always dissolved last. Each 5 mg of nitroBT is dissolved in 40 &#181;L mix (20 &#181;L ethanol + 20 &#181;L dimethylformamide). All volumes are per 1 mL of 18% PVA solution, which can be used to stain ~2 tissue sections or ~4 cell preparations on glass slides. NitroBT, NAD+, NADP+, ADP and substrate solutions should be made freshly. NaN3, MgCl2 and (m)PMS solutions can be stored at 4 &#176;C for one month. 18% PVA dissolved in phosphate buffer can be stored at 60 &#176;C for 2 weeks. The pH of the 18% PVA solution can be set using different compositions of the 0.1 M potassium dihydrogen phosphate (KH2PO4) and 0.1 M disodium hydrogen phosphate (NA2HPO4) buffers. The shown incubation periods provide good starting points, but the optimal incubation periods depend on the species, the tissue type and the preservation of the tissue. In general, cell preparations should be incubated longer than tissue sections to obtain a similar level of formazan production.
Wild-type isocitrate dehydrogenase 1 and 2 (IDH1/2) catalyze the conversion of isocitrate to &#945;-ketoglutarate (&#945;KG) with concomitant reduction of NADP+ to NADPH in the cytoplasm and mitochondria, respectively. These enzymes have recently attracted interest because IDH1/2 mutations occur in various types of human cancer, including primary brain cancer (glioblastoma) and colorectal cancer, and offer a unique opportunity to demonstrate the possibilities of metabolic mapping. IDH1 mutations disable IDH1 wild-type enzyme activity and also induce a neo-enzymatic activity that leads to the production and subsequent accumulation of the oncometabolite D-2-hydroxyglutarate (D-2HG),11 which is discussed elsewhere in detail.12 Metabolic mapping experiments showed that NADP+-dependent IDH1/2 activity was significantly lower in colorectal carcinoma cells with a knocked-in heterozygous IDH1 mutation than in colorectal carcinoma cells that were IDH1 wild-type (Figure 1A). This difference was quantified by image analysis of monochromatic light photomicrographs (Figure 1B).12
Another illustrating example of metabolic mapping experiments is shown in Figure 2, which is an image of a cryostat section of mouse brain that was injected with human glioblastoma cells. IDH1/2 is the most important NADPH provider in human brain and its activity is upregulated in wild-type IDH1/2 glioblastoma, whereas the NADPH production capacity of IDH1/2 is much lower in the brains of rodents.11 Figure 2A shows the difference between the high NADP+-dependent IDH1/2 activity in human glioblastoma cells and the low NADP+-dependent IDH1/2 activity in healthy mouse brain. IDH1/2 activity is quantified as &#181;mol NADPH production per mL of tissue per minute in Figure 2B.
The enzymatic reaction of IDH1/2 is relatively straightforward, but lactate dehydrogenase (LDH) is a more complicated enzyme complex. LDH converts the reversible conversion from lactate to pyruvate with concomitant reduction of NAD+ to NADH or vice versa. The availability of lactate and pyruvate and the composition of the LDH enzyme complex dictates whether lactate is primarily converted to pyruvate (which is catalyzed by LDH-B and yields NADH) or whether pyruvate is primarily converted to lactate (which is catalyzed by LDH-A and consumes NADH).13 Metabolic mapping experiments are able to visualize the activity of the LDH-B reaction, but they are &#34;blind&#34; to the activity of the LDH-A reaction because NADH production does not occur and as a consequence nitroBT is not reduced to formazan. Figure 3 shows the NAD+-dependent LDH activity (i.e. the conversion of lactate into pyruvate) in human brain sections in the absence of substrate (Figure 3A), in the presence of a high concentration of lactate (Figure 3B), in the presence of&#160;6 mM lactate (Figure 3C) and in the presence of a low concentration of lactate and a higher concentration of pyruvate (Figure 3D). The results of these experiments illustrate metabolic mapping adequately captures that the NAD+-dependent LDH activity in tissue sections depends on the availability of lactate and pyruvate in the reaction medium.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56843/56843fig1.jpg" />
Figure 1. NADP+-dependent IDH1/2 activity of human colorectal carcinoma cells. (A) Representative monochromatic light photomicrographs of HCT116 human colorectal carcinoma cells after staining for NADP+-dependent IDH1/2 activity against 1-10 mM isocitrate and 0.8 mM NADP+. Scale bars = 50 &#181;m. (B) A quantification of the absorbance of blue formazan produced from nitroBT by NADP+-dependent IDH1/2 activity per cell is shown with the use of monochromatic light in (A) and image analysis. Abbreviations: IDH1WT/WT, isocitrate dehydrogenase 1 wild-type;&#160;IDH1WT/MUT, isocitrate dehydrogenase 1 mutated.&#160;<a href="https://cloudflare.jove.com/files/ftp_upload/56843/56843fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56843/56843fig2.jpg" />
Figure 2. NADP+-dependent IDH1/2 activity of human glioblastoma samples. (A) Representative photomicrograph of a mouse brain section containing healthy mouse brain (h) and a human glioblastoma tumor xenograft (t) after staining for NADP+-dependent IDH1/2 activity in the presence of 0.8 mM NADP+ and 2 mM isocitrate. Scale bar = 200 &#181;m (B) Enzyme activity quantification of areas of mouse brain (h) and human glioblastoma xenograft (t) shown in (A) using the law of Lambert-Beer. Error bars represent standard deviations. <a href="https://cloudflare.jove.com/files/ftp_upload/56843/56843fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56843/56843fig3.jpg" />
Figure 3. NAD+-dependent lactate dehydrogenase (LDH) activity of human brain samples. Representative photomicrographs of serial sections of human brain samples after staining for NAD+-dependent LDH activity (A) against control conditions (in the absence substrate and pyruvate, 3 mM NAD+ only) (B) against 150 mM lactate in the absence of pyruvate (C) against 6 mM lactate in the absence of pyruvate and (D) against 6 mM lactate in the presence of 18 mM pyruvate, the competitive product inhibitor of the lactate-to-pyruvate reaction. Scale bar = 200 &#181;m. <a href="https://cloudflare.jove.com/files/ftp_upload/56843/56843fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Enzyme:</td>
			<td>G6PDH</td>
			<td>LDH</td>
			<td>SDH</td>
			<td>MDH</td>
			<td>IDH1</td>
			<td>IDH2</td>
			<td>IDH3</td>
			<td>6PGD</td>
			<td>GDH</td>
		</tr>
		<tr>
			<td>PVA 18% in 100 mM phosphate buffer</td>
			<td>1 mL; pH = 7.4</td>
			<td>1 mL;&#160; pH = 7.4</td>
			<td>1 mL;&#160; pH = 8.0</td>
			<td>1 mL; pH = 7.4</td>
			<td>1 mL; pH = 7.4</td>
			<td>1 mL; pH = 7.4</td>
			<td>1 mL; pH = 7.4</td>
			<td>1 mL; pH = 8.0</td>
			<td>1 mL; pH = 8.0</td>
		</tr>
		<tr>
			<td>5 mM NitroBT</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
			<td>5 mg/40 &#181;L mix</td>
		</tr>
		<tr>
			<td>5 mM NaN3&#160;</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
		</tr>
		<tr>
			<td>5 mM MgCl2&#160;</td>
			<td>10 &#181;L</td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td></td>
		</tr>
		<tr>
			<td>3 mM NAD+</td>
			<td></td>
			<td>10 &#181;L</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
			<td></td>
			<td>10 &#181;L</td>
		</tr>
		<tr>
			<td>0.8 mM NADP</td>
			<td>10 &#181;L</td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td></td>
			<td>10 &#181;L</td>
			<td></td>
		</tr>
		<tr>
			<td>2 mM ADP</td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
		</tr>
		<tr>
			<td>substrate</td>
			<td>10 mM glucose-6-phosphate</td>
			<td>150 mM lactate</td>
			<td>50 mM succinate</td>
			<td>100 mM malic acid</td>
			<td>2 mM isocitrate&#160;</td>
			<td>2 mM isocitrate</td>
			<td>2 mM isocitrate</td>
			<td>10 mM 6-phospho-galactonate</td>
			<td>10 mM glutamic acid</td>
		</tr>
		<tr>
			<td>0.32 mM mPMS&#160;</td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td></td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
			<td></td>
		</tr>
		<tr>
			<td>0.2 mM PMS&#160;</td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
			<td></td>
			<td></td>
			<td>10 &#181;L</td>
			<td>10 &#181;L</td>
			<td></td>
			<td>10 &#181;L</td>
		</tr>
		<tr>
			<td>Incubation duration</td>
			<td>5 min</td>
			<td>30 min</td>
			<td>60 min</td>
			<td>60 min</td>
			<td>30 min</td>
			<td>30 min</td>
			<td>30 min</td>
			<td>10 min</td>
			<td>30 min</td>
		</tr>
	</tbody>
</table>

Table 1. Schematic overview of the metabolic mapping protocol for dehydrogenases.&#160;&#160;Abbreviations: ADP, adenosine diphosphate; (m)PMS (methoxy)-5-methylphenazinium methyl sulfate; nitroBT, nitro tetrazolium blue chloride; G6PDH, glucose-6-phosphate dehydrogenase; LDH, lactate dehydrogenase; SDH, succinate dehydrogenase; MDH, malate dehydrogenase; IDH, isocitrate dehydrogenase; 6PGD, 6-phosphogluconate dehydrogenase; GDH, glutamate dehydrogenase.

Watch this Scientific Journal Video about Metabolic Mapping: Quantitative Enzyme Cytochemistry and Histochemistry to Determine the Activity of Dehydrogenases in Cells and Tissues at JoVE.com

Metabolic Mapping: Quantitative Enzyme Cytochemistry and Histochemistry to Determine the Activity of Dehydrogenases in Cells and Tissues

Here, we present a protocol that can be used to microscopically visualize and quantify activity of dehydrogenases in cells and tissues and its kinetics, function and subcellular localization.

Altered cellular metabolism is a hallmark of many diseases, including cancer, cardiovascular diseases and infection. The metabolic motor units of cells ...

metabolic-mapping-quantitative-enzyme-cytochemistry-histochemistry-to

Research

JoVE Journal

Immunology and Infection

11.6K Views.  University of Amsterdam, Amsterdam, The Netherlands. Here, we present a protocol that can be used to microscopically visualize and quantify activity of dehydrogenases in cells and tissues and its kinetics, function and subcellular localization.

Video: Metabolic Mapping: Quantitative Enzyme Cytochemistry and Histochemistry to Determine the Activity of Dehydrogenases in Cells and Tissues

Seven Steps to Stellate Cells

Hepatic stellate cells are liver-resident cells of star-like morphology and are located in the space of Disse between liver sinusoidal endothelial cells and hepatocytes1,2. Stellate cells are derived from bone marrow precursors and store up to 80% of the total body vitamin A1, 2. Upon activation, stellate cells differentiate into myofibroblasts to produce extracellular matrix, thus contributing to liver fibrosis3. Based on their ability to contract, myofibroblastic stellate cells can regulate the vascular tone associated with portal hypertension4. Recently, we demonstrated that hepatic stellate cells are potent antigen presenting cells and can activate NKT cells as well as conventional T lymphocytes5. 

 Here we present a method for the efficient preparation of hepatic stellate cells from mouse liver. Due to their perisinusoidal localization, the isolation of hepatic stellate cells is a multi-step process. In order to render stellate cells accessible to isolation from the space of Disse, mouse livers are perfused in situ with the digestive enzymes Pronase E and Collagenase P. Following perfusion, the liver tissue is subjected to additional enzymatic treatment with Pronase E and Collagenase P in vitro. Subsequently, the method takes advantage of the massive amount of vitamin A-storing lipid droplets in hepatic stellate cells. This feature allows the separation of stellate cells from other hepatic cell types by centrifugation on an 8% Nycodenz gradient. The protocol described here yields a highly pure and homogenous population of stellate cells. Purity of preparations can be assessed by staining for the marker molecule glial fibrillary acidic protein (GFAP), prior to analysis by fluorescence microscopy or flow cytometry. Further, light microscopy reveals the unique appearance of star-shaped hepatic stellate cells that harbor high amounts of lipid droplets. 


 Taken together, we present a detailed protocol for the efficient isolation of hepatic stellate cells, including representative images of their morphological appearance and GFAP expression that help to define the stellate cell entity.

Here we describe a method for the isolation of hepatic stellate cells from mouse liver. For stellate cell purification, mouse livers are digested in situ and in vitro by pronase-collagenase treatment prior to density gradient centrifugation. This technique yields highly pure hepatic stellate cells.

Hepatic stellate cells are liver-resident  cells of star-like morphology and are located in the space of Disse between  liver sinusoidal endothelial cells ...

Following Cell-fate in E. coli After Infection by Phage Lambda

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the simultaneous infection of the cell by a number of phages, one of two pathways is chosen: lytic (virulent) or lysogenic (dormant)3,4. We recently developed a method for fluorescently labeling individual phages, and were able to examine the post-infection decision in real-time under the microscope, at the level of individual phages and cells5. Here, we describe the full procedure for performing the infection experiments described in our earlier work5. This includes the creation of fluorescent phages, infection of the cells, imaging under the microscope and data analysis. The fluorescent phage is a "hybrid", co-expressing wild- type and YFP-fusion versions of the capsid gpD protein. A crude phage lysate is first obtained by inducing a lysogen of the gpD-EYFP (Enhanced Yellow Fluorescent Protein) phage, harboring a plasmid expressing wild type gpD. A series of purification steps are then performed, followed by DAPI-labeling and imaging under the microscope. This is done in order to verify the uniformity, DNA packaging efficiency, fluorescence signal and structural stability of the phage stock. The initial adsorption of phages to bacteria is performed on ice, then followed by a short incubation at 35&deg;C to trigger viral DNA injection6. The phage/bacteria mixture is then moved to the surface of a thin nutrient agar slab, covered with a coverslip and imaged under an epifluorescence microscope. The post-infection process is followed for 4 hr, at 10 min interval. Multiple stage positions are tracked such that ~100 cell infections can be traced in a single experiment. At each position and time point, images are acquired in the phase-contrast and red and green fluorescent channels. The phase-contrast image is used later for automated cell recognition while the fluorescent channels are used to characterize the infection outcome: production of new fluorescent phages (green) followed by cell lysis, or expression of lysogeny factors (red) followed by resumed cell growth and division. The acquired time-lapse movies are processed using a combination of manual and automated methods. Data analysis results in the identification of infection parameters for each infection event (e.g. number and positions of infecting phages) as well as infection outcome (lysis/lysogeny). Additional parameters can be extracted if desired.

Following Cell-fate in E. coli After Infection by Phage Lambda

This article describes the procedure for preparing a fluorescently-labeled version of bacteriophage lambda, infection of E. coli bacteria, following the infection outcome under the microscope, and analysis of infection results.

The system comprising bacteriophage (phage) lambda and the bacterium E. coli has long served as a paradigm for cell-fate determination1,2. Following the ...

Assessing Hepatic Metabolic Changes During Progressive Colonization of Germ-free Mouse by 1H NMR Spectroscopy

It is well known that gut bacteria contribute significantly to the host homeostasis, providing a range of benefits such as immune protection and vitamin synthesis. They also supply the host with a considerable amount of nutrients, making this ecosystem an essential metabolic organ. In the context of increasing evidence of the link between the gut flora and the metabolic syndrome, understanding the metabolic interaction between the host and its gut microbiota is becoming an important challenge of modern biology.1-4
 Colonization (also referred to as normalization process) designates the establishment of micro-organisms in a former germ-free animal. While it is a natural process occurring at birth, it is also used in adult germ-free animals to control the gut floral ecosystem and further determine its impact on the host metabolism. A common procedure to control the colonization process is to use the gavage method with a single or a mixture of micro-organisms. This method results in a very quick colonization and presents the disadvantage of being extremely stressful5. It is therefore useful to minimize the stress and to obtain a slower colonization process to observe gradually the impact of bacterial establishment on the host metabolism.
 In this manuscript, we describe a procedure to assess the modification of hepatic metabolism during a gradual colonization process using a non-destructive metabolic profiling technique. We propose to monitor gut microbial colonization by assessing the gut microbial metabolic activity reflected by the urinary excretion of microbial co-metabolites by 1H NMR-based metabolic profiling. This allows an appreciation of the stability of gut microbial activity beyond the stable establishment of the gut microbial ecosystem usually assessed by monitoring fecal bacteria by DGGE (denaturing gradient gel electrophoresis).6 The colonization takes place in a conventional open environment and is initiated by a dirty litter soiled by conventional animals, which will serve as controls. Rodents being coprophagous animals, this ensures a homogenous colonization as previously described.7
 Hepatic metabolic profiling is measured directly from an intact liver biopsy using 1H High Resolution Magic Angle Spinning NMR spectroscopy. This semi-quantitative technique offers a quick way to assess, without damaging the cell structure, the major metabolites such as triglycerides, glucose and glycogen in order to further estimate the complex interaction between the colonization process and the hepatic metabolism7-10. This method can also be applied to any tissue biopsy11,12.

Assessing Hepatic Metabolic Changes During Progressive Colonization of Germ-free Mouse by 1H NMR Spectroscopy

A progressive colonization procedure is described to further assess its impact on the host hepatic metabolism. Colonization is monitored non invasively by evaluating the urinary excretion of microbial co-metabolites by NMR-based metabolic profiling while hepatic metabolism is assessed by High Resolution Magic Angle Spinning (HR MAS) NMR profiling of intact biopsy.

It is well known that gut bacteria contribute significantly to the host homeostasis, providing a range of benefits such as immune protection and vitamin ...

Quantitative In vitro Assay to Measure Neutrophil Adhesion to Activated Primary Human Microvascular Endothelial Cells under Static Conditions

The vascular endothelium plays an integral part in the inflammatory response. During the acute phase of inflammation, endothelial cells (ECs) are activated by host mediators or directly by conserved microbial components or host-derived danger molecules. Activated ECs express cytokines, chemokines and adhesion molecules that mobilize, activate and retain leukocytes at the site of infection or injury. Neutrophils are the first leukocytes to arrive, and adhere to the endothelium through a variety of adhesion molecules present on the surfaces of both cells. The main functions of neutrophils are to directly eliminate microbial threats, promote the recruitment of other leukocytes through the release of additional factors, and initiate wound repair. Therefore, their recruitment and attachment to the endothelium is a critical step in the initiation of the inflammatory response. In this report, we describe an in vitro neutrophil adhesion assay using calcein AM-labeled primary human neutrophils to quantitate the extent of microvascular endothelial cell activation under static conditions. This method has the additional advantage that the same samples quantitated by fluorescence spectrophotometry can also be visualized directly using fluorescence microscopy for a more qualitative assessment of neutrophil binding.

Quantitative In vitro Assay to Measure Neutrophil Adhesion to Activated Primary Human Microvascular Endothelial Cells under Static Conditions

Neutrophil adherence to the activated endothelium at sites of infection is an integral component of the host's inflammatory response. Described in this report is a neutrophil binding assay that allows for the in vitro quantitation of primary human neutrophil binding to endothelial cells activated by inflammatory mediators under static conditions.

The vascular  endothelium plays an integral part in the inflammatory response. During the  acute phase of inflammation, endothelial cells (ECs) are ...

Methods to Inhibit Bacterial Pyomelanin Production and Determine the Corresponding Increase in Sensitivity to Oxidative Stress

Pyomelanin is an extracellular red-brown pigment produced by several bacterial and fungal species. This pigment is derived from the tyrosine catabolism pathway and contributes to increased oxidative stress resistance. Pyomelanin production in Pseudomonas aeruginosa is reduced in a dose dependent manner through treatment with 2-[2-nitro-4-(trifluoromethyl)benzoyl]-1,3-cyclohexanedione (NTBC). We describe a titration method using multiple concentrations of NTBC to determine the concentration of drug that will reduce or abolish pyomelanin production in bacteria. The titration method has an easily quantifiable outcome, a visible reduction in pigment production with increasing drug concentrations. We also describe a microtiter plate method to assay antibiotic minimum inhibitory concentration (MIC) in bacteria. This method uses a minimum of resources and can easily be scaled up to test multiple antibiotics in one microtiter plate for one strain of bacteria. The MIC assay can be adapted to test the affects of non-antibiotic compounds on bacterial growth at specific concentrations. Finally, we describe a method for testing bacterial sensitivity to oxidative stress by incorporating H2O2 into agar plates and spotting multiple dilutions of bacteria onto the plates. Sensitivity to oxidative stress is indicated by reductions in colony number and size for the different dilutions on plates containing H2O2 compared to a no H2O2 control. The oxidative stress spot plate assay uses a minimum of resources and low concentrations of H2O2. Importantly, it also has good reproducibility. This spot plate assay could be adapted to test bacterial sensitivity to various compounds by incorporating the compounds in agar plates and characterizing the resulting bacterial growth.

Bacterial pyomelanin production results in increased resistance to oxidative stress and virulence. We report on techniques that can be used to determine inhibition of pyomelanin production and assay the resulting increase in sensitivity to oxidative stress in bacteria, as well as determine antibiotic minimum inhibitory concentration (MIC).

Pyomelanin is an extracellular red-brown pigment produced by several bacterial and fungal species. This pigment is derived from the tyrosine catabolism ...

Neutrophil Isolation and Analysis to Determine their Role in Lymphoma Cell Sensitivity to Therapeutic Agents

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of inflammation. They are key players in the innate immune system and play major roles in cancer biology. Neutrophils have been proposed as key mediators of malignant transformation, tumor progression, angiogenesis and in the modulation of the antitumor immunity; through their release of soluble factors or their interaction with tumor cells. To characterize the specific functions of neutrophils, a fast and reliable method is coveted for in vitro isolation of neutrophils from human blood. Here, a density gradient separation method is demonstrated to isolate neutrophils as well as mononuclear cells from the blood. The procedure consists of layering the density gradient solution such as Ficoll carefully above the diluted blood obtained from patients diagnosed with chronic lymphocytic leukemia (CLL), followed by centrifugation, isolation of mononuclear layer, separation of neutrophils from RBCsby dextran then lysis of residual erythrocytes. This method has been shown to isolate neutrophils &#8805; 90 % pure. To mimic the tumor microenvironment, 3-dimensional (3D) experiments were performed using basement membrane matrix such as Matrigel. Given the short half-life of neutrophils in vitro, 3D experiments with fresh human neutrophils cannot be performed. For this reason promyelocytic HL60 cells are differentiated along the granulocytic pathway using the differentiation inducers dimethyl sulfoxide (DMSO) and retinoic acid (RA). The aim of our experiments is to study the role of neutrophils on the sensitivity of lymphoma cells to anti-lymphoma agents. However these methods can be generalized to study the interactions of neutrophils or neutrophil-like cells with a large range of cell types in different situations.

Neutrophils are the most abundant type of white blood cells. The isolation of neutrophils from human blood by density gradient separation method and the differentiation of human promyelocytic (HL60) cells along the granulocytic pathway are described here; to test their role on sensitivity of lymphoma cells to anti-lymphoma agents.

Neutrophils are the most abundant (40% to 75%) type of white blood cells and among the first inflammatory cells to migrate towards the site of ...

Optimization of a Quantitative Micro-neutralization Assay

The micro-neutralization (MN) assay is a standard technique for measuring the infectivity of the influenza virus and the inhibition of virus replication. In this study, we present the protocol of an imaging-based MN assay to quantify the true antigenic relationships between viruses. Unlike typical plaque reduction assays that rely on visible plaques, this assay quantitates the entire infected cell population of each well. The protocol matches the virus type or subtype with the selection of cell lines to achieve maximum infectivity, which enhances sample contrast during imaging and image processing. The introduction of quantitative titration defines the amount of input viruses of neutralization and enables the results from different experiments to be comparable. The imaging setup with a flatbed scanner and free downloadable software makes the approach high throughput, cost effective, user friendly, and easy to deploy in most laboratories. Our study demonstrates that the improved MN assay works well with the current circulating influenza A(H1N1)pdm09, A(H3N2), and B viruses, without being significantly influenced by amino acid substitutions in the neuraminidase (NA) of A(H3N2) viruses. It is particularly useful for the characterization of viruses that either grow to low HA titer and/or undergo an abortive infection resulting in an inability to form plaques in cultured cells.

This study describes an imaging-based micro-neutralization assay to analyze the antigenic relationships between viruses. The protocol employs a flatbed scanner and has four steps, including titration, titration quantitation, neutralization, and neutralization quantitation. The assay works well with current circulating influenza A(H1N1)pdm09, A(H3N2), and B viruses.

The micro-neutralization (MN) assay is a standard technique for measuring the infectivity of the influenza virus and the inhibition of virus replication. ...

In Situ MHC-tetramer Staining and Quantitative Analysis to Determine the Location, Abundance, and Phenotype of Antigen-specific CD8 T Cells in Tissues

T cells are critical to many immunological processes, including detecting and eliminating virus-infected cells, preventing autoimmunity, assisting in B-cell and plasma-cell production of antibodies, and detecting and eliminating cancer cells. The development of MHC-tetramer staining of antigen-specific T cells analyzed by flow cytometry has revolutionized our ability to study and understand the immunobiology of T cells. While extremely useful for determining the quantity and phenotype of antigen-specific T cells, flow cytometry cannot determine the spatial localization of antigen-specific T cells to other cells and structures in tissues, and current disaggregation techniques to extract the T cells needed for flow cytometry have limited effectiveness in non-lymphoid tissues. In situ MHC-tetramer staining (IST) is a technique to visualize T cells that are specific for antigens of interest in tissues. In combination with immunohistochemistry (IHC), IST can determine the abundance, location, and phenotype of antigen-specific CD8 and CD4 T cells in tissues. Here, we describe a protocol to stain and enumerate antigen-specific CD8 T cells, with specific phenotypes located within specific tissue compartments. These procedures are the same that we used in our recent publication by Li et al., entitled &#34;Simian Immunodeficiency Virus-Producing Cells in Follicles Are Partially Suppressed by CD8+ Cells In Vivo.&#34; The methods described are broadly applicable because they can be used to localize, phenotype, and quantify essentially any antigen-specific CD8 T cell for which MHC tetramers are available, in any tissue.

In Situ MHC-tetramer Staining and Quantitative Analysis to Determine the Location, Abundance, and Phenotype of Antigen-specific CD8 T Cells in Tissues

Here, we describe a method that combines in situ MHC-tetramer staining with immunohistochemistry to determine localization, phenotype, and quantity of antigen-specific T cells in tissues. This protocol is used to determine the spatial and phenotypic characteristics of antigen-specific CD8 T cells relative to other cell type and structures in tissues.

T cells are critical to many immunological processes, including detecting and eliminating virus-infected cells, preventing autoimmunity, assisting in ...

Overexpression and Purification of Human Cis-prenyltransferase in Escherichia coli

Prenyltransferases (PT) are a group of enzymes that catalyze chain elongation of allylic diphosphate using isopentenyl diphosphate (IPP) via multiple condensation reactions. DHDDS (dehydrodolichyl diphosphate synthase) is a eukaryotic long-chain cis-PT (forming cis double bonds from the condensation reaction) that catalyzes chain elongation of farnesyl diphosphate (FPP, an allylic diphosphate) via multiple condensations with isopentenyl diphosphate (IPP). DHDDS is of biomedical importance, as a non-conservative mutation (K42E) in the enzyme results in retinitis pigmentosa, ultimately leading to blindness. Therefore, the present protocol was developed in order to acquire large quantities of purified DHDDS, suitable for mechanistic studies. Here, the usage of protein fusion, optimized culture conditions and codon-optimization were used to allow the overexpression and purification of functionally active human DHDDS in E. coli. The described protocol is simple, cost-effective and time sparing. The homology of cis-PT among different species suggests that this protocol may be applied for other eukaryotic cis-PT as well, such as those involved in natural rubber synthesis.

Overexpression and Purification of Human Cis-prenyltransferase in Escherichia coli

A simple protocol for overexpression and purification of codon-optimized, human cis-prenyltransferase, under non-denaturing conditions, from Escherichia coli, is described, along with an enzymatic activity assay. This protocol can be generalized for production of other cis- prenyltransferase proteins in quantity and quality suitable for mechanistic studies.

Prenyltransferases (PT) are a group of enzymes that catalyze chain elongation of allylic diphosphate using isopentenyl diphosphate (IPP) via multiple ...

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro

Natural killer (NK) cells belong to the innate immune system and are a first-line anti-cancer immune defense; however, they are suppressed in the tumor microenvironment and the underlying mechanism is still largely unknown. The lack of a consistent and reliable source of NK cells limits the research progress of NK cell immunity. Here, we report an in vitro system that can provide high quality and quantity of bone marrow-derived murine NK cells under a feeder-free condition. More importantly, we also demonstrate that siRNA-mediated gene silencing successfully inhibits the E4bp4-dependent NK cell maturation by using this system. Thus, this novel in vitro NK cell differentiating system is a biomaterial solution for immunity research.

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro

Here, we present a protocol to mass-produce gene-silencing murine NK cells by using a feeder-free differentiation system for mechanistic study in vitro and in vivo.

Natural killer (NK) cells belong to the innate immune system and are a first-line anti-cancer immune defense; however, they are suppressed in the tumor ...

Metabolic Mapping: Quantitative Enzyme Cytochemistry and Histochemistry to Determine the Activity of Dehydrogenases in Cells and Tissues

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Chapters in this video

Seven Steps to Stellate Cells

Following Cell-fate in E. coli After Infection by Phage Lambda

Assessing Hepatic Metabolic Changes During Progressive Colonization of Germ-free Mouse by ¹H NMR Spectroscopy

Quantitative In vitro Assay to Measure Neutrophil Adhesion to Activated Primary Human Microvascular Endothelial Cells under Static Conditions

Methods to Inhibit Bacterial Pyomelanin Production and Determine the Corresponding Increase in Sensitivity to Oxidative Stress

Neutrophil Isolation and Analysis to Determine their Role in Lymphoma Cell Sensitivity to Therapeutic Agents

Optimization of a Quantitative Micro-neutralization Assay

In Situ MHC-tetramer Staining and Quantitative Analysis to Determine the Location, Abundance, and Phenotype of Antigen-specific CD8 T Cells in Tissues

Overexpression and Purification of Human Cis-prenyltransferase in Escherichia coli

A Novel Feeder-free System for Mass Production of Murine Natural Killer Cells In Vitro