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 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's activity. The formazan's absorbance is therefore a direct measure of the dehydrogenase'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.
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.
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
2. Enzyme Histo/Cytochemistry for Soluble Dehydrogenases
3. Image Acquisition
4. Image 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 °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 µL mix (20 µL ethanol + 20 µL 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 °C for one month. 18% PVA dissolved in phosphate buffer can be stored at 60 °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 α-ketoglutarate (αKG) 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 µmol 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 "blind" 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 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.
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 µm. (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; IDH1WT/MUT, isocitrate dehydrogenase 1 mutated. Please click here to view a larger version of this figure.
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 µm (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. Please click here to view a larger version of this figure.
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 µm. Please click here to view a larger version of this figure.
Enzyme: | G6PDH | LDH | SDH | MDH | IDH1 | IDH2 | IDH3 | 6PGD | GDH |
PVA 18% in 100 mM phosphate buffer | 1 mL; pH = 7.4 | 1 mL; pH = 7.4 | 1 mL; pH = 8.0 | 1 mL; pH = 7.4 | 1 mL; pH = 7.4 | 1 mL; pH = 7.4 | 1 mL; pH = 7.4 | 1 mL; pH = 8.0 | 1 mL; pH = 8.0 |
5 mM NitroBT | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix | 5 mg/40 µL mix |
5 mM NaN3 | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL |
5 mM MgCl2 | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | |||
3 mM NAD+ | 10 µL | 10 µL | 10 µL | ||||||
0.8 mM NADP | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | ||||
2 mM ADP | 10 µL | ||||||||
substrate | 10 mM glucose-6-phosphate | 150 mM lactate | 50 mM succinate | 100 mM malic acid | 2 mM isocitrate | 2 mM isocitrate | 2 mM isocitrate | 10 mM 6-phospho-galactonate | 10 mM glutamic acid |
0.32 mM mPMS | 10 µL | 10 µL | 10 µL | 10 µL | 10 µL | ||||
0.2 mM PMS | 10 µL | 10 µL | 10 µL | 10 µL | |||||
Incubation duration | 5 min | 30 min | 60 min | 60 min | 30 min | 30 min | 30 min | 10 min | 30 min |
Table 1. Schematic overview of the metabolic mapping protocol for dehydrogenases. 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.
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.
The authors have nothing to disclose.
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.
Adenosine 5′-diphosphate sodium salt | Sigma-Aldrich | A2754 | |
D-Glucose 6-phosphate sodium salt | Sigma-Aldrich | G7879 | |
Disodium hydrogen phosphate (NA2HPO4) | Merck Millipore | 106559 | |
di-Sodium succinate | Sigma-Aldrich | W327700 | |
DL-Isocitric acid trisodium salt hydrate | Sigma-Aldrich | I1252 | |
D-Malic acid | Sigma-Aldrich | 2300 | |
Glycerol Gelatin | Sigma-Aldrich | GG1 | |
L-Glutamic acid | Sigma-Aldrich | G1251 | |
Lithium 6-phospho-D-galactonate | Sigma-Aldrich | 55962 | |
Magnesium chloride | Sigma-Aldrich | M8266 | |
methoxy-Phenazine methosulfate | Serva | 28966.01 | |
Nicotinamide adenine dinucleotide | Sigma-Aldrich | N0505 | |
Nicotinamide adenine dinucleotide phosphate | Sigma-Aldrich | NADP-RO | |
Nitrotetrazolium Blue chloride | Sigma-Aldrich | N6876 | |
Phenazine methosulfate | Serva | 32030.01 | |
Polyvinyl alcohol, fully hydrolyzed | Sigma-Aldrich | P1763 | |
Potassium dihydrogen phosphate (KH2PO4) | Merck Millipore | 104873 | |
Sodium azide | Sigma-Aldrich | S2002 | |
Sodium L-lactate | Sigma-Aldrich | L7022 | |
Bandpass filter | Edmund optics | https://www.edmundoptics.de/optics/optical-filters/bandpass-filters/Hard-Coated-OD-4-10nm-Bandpass-Filters/ | |
Grey-step wedge | Stouffer Industries | http://www.stouffer.net/TransPage.htm |