Assessment of Nutrient Uptake Metabolite Production and Enzyme Activity

1. MICROFLUOROMETRIC TECHNIQUES

The ability to measure the uptake and production of substrates by single embryos is an important tool for understanding how embryos control metabolism and embryo production. Little is known about the control of metabolism in the mammalian embryo compared to somatic cells or embryos from marine species or Xenopus. One of the main reasons for this is that the small amounts of material in the early embryo make measurement technically difficult. In recent years microfluorometric techniques have been developed that are capable of quantitatively analyzing the nutrient uptake and metabolite release from single cells. These techniques are based on miniaturizations of conventional methods of enzymatic analysis and can therefore be adapted to measure a great variety of metabolites (1-3) (Figure 8.1). An advantage of these procedures over the radiolabel procedures described in Chapter 9 is that they are totally noninvasive. Thus, microfluorometric techniques have the potential to be used as viability assays for mammalian embryos (2).

In addition, microfluorometry can be used to measure the enzyme activity in a single cell. It is possible to examine the activity and kinetics of a specific enzyme from a single oocyte or embryo using these procedures. In this chapter we provide details about how to use ultra-microfluriometry to measure nutrient uptakes and enzyme activities in mammalian embryos. Additionally, information is provided on how to calculate the kinetic properties of the enzymes and how to identify the isoforms present using electrophoresis.

2. ULTRAMICROFLUORIMETRY

Microfluorometric techniques developed in the 1980s are capable of quantitatively analyzing the nutrient uptake and metabolite release from single cells (4). These techniques are miniaturizations of conventional fluorometric methods of enzymatic analysis. Rather than occurring in cuvettes, the assays are scaled down with the use of micropipettes to measure picomole/femtomole levels of substrates.

Uptake

Production

Glucost Pyruvate Amino Acids

Other Sugars Oxygen

Uptake And Production From Embryo
Growth Factors

Lactate

Ammonium

Enzymes

Defined Medium

Embryo

Figure 8.1. Diagram of how substrate uptake and production by embryos can be measured using ultramicrofluorimetry. Individual embryos are placed in known volumes of defined medium. The uptake of a substrate can be determined by measuring its rate of disappearance from the medium, whereas the production of a metabolite can be measured by its rate of appearance in the medium.

Fluorometric assays are based on the generation or consumption of the reduced pyridine nucleotides, NADH and NADPH, in coupled enzymatic reactions. These nucleotides fluoresce when excited with light at 340 nm, whereas the oxidized forms, NAD+ and NADP+, do not. Thus, the reaction:

may be followed by monitoring a drop in fluorescence, or optical density at 340 nm, as NADH is converted to NAD+. Under specific conditions the levels of change in fluorescence are proportional to the amount of substrate consumed in the reaction, and therefore the amounts can be calibrated using standard curves. Reactions are conventionally carried out in cuvettes and the fluorescence or absorbance measured by a fluorimeter or spectrophotometer. However, to study single-cell biochemistry, reactions are scaled down to occur in nanoliter volumes so that substrate levels in the picomole and femtomole range can be measured.

It is possible to measure any substrate that can be linked to the reaction that contains pyridine nucleotides or another fluorescent tag (Figure 8.1).

3. EQUIPMENT REQUIREMENTS

Assays for substrate levels and enzyme activity rely on quantiating the fluorescence of the reduced forms of the pyridine nucleotides (NADH, NADPH) under UV light. Therefore, the essentials of the equipment for this type of analysis are excitation of a sample with a mercury lamp so that excitation wavelengths are in the UV range (340 nm), a diaphragm to limit the excitation and emission to the drops to be analyzed, and quantification of the low emission levels from the sample. This is best achieved by the use of a photometer attachment with software that can convert the levels of emission light into a numerical value. The photometer scale is linear and is expressed in arbitrary units that can be calibrated using standard curves. Additionally, for the measurement of enzyme activity the use of an optical switch (as opposed to the use of shutters) is preferable because it enables real-time continuous kinetic measurement of enzyme activity.

Lactate dehydrogenase Pyruvate + NADH + H+ ^ Lactate + NAD

Additional equipment to the fluorimetry measurement system that are required are a micromanipulator and stereo microscope for manipulation of micropipettes, a warm stage for embryos, and a warm stage on fluorimetry equipment for measurement of enzyme activities.

4. MICROPIPETTES

For analysis of substrate uptakes, metabolite production, or enzyme activity by embryos, the conventional assays are scaled down to occur in submicroliter volumes. The sub-microliter volumes are manipulated by specially constructed constriction micropipettes. These pipettes are made from borosilicate glass capillaries (o.d. 1.0 mm, i.d. 0.8 mm) pulled over a flame to produce an inner diameter of 50-100 mm. The tubing is snapped in half and a small hook made on each end. Using a microforge, a constriction is made in the glass by placing the heated filament close to the glass capillary. A small weight (paperclip) is then placed onto the end of the hook of the capillary, and the tip is made by again heating the glass with the microforge filament. As the glass heats up, the weight will result in the glass pulling to a tip. The tip can then be broken using a pair of watchmaker's forceps. The size of the tip and constriction will control the speed and accuracy at which the pipette can be filled. The glass pipettes are mounted in 16-gauge stainless-steel tubing and sealed using sealing wax. Filling and expelling fluids from the pipettes is achieved using an air-filled syringe attached to the pipette via thin tubing. Before use, the micropipettes are siliconized to make manipulation of fluids easier (Figure 8.2). The volume of the pipettes between the tip and the constriction is calibrated using tritiated water. The pipettes are then held in a micromanipulator and the volumes manipulated under a microscope. Using this procedure, it is possible to accurately pipette volumes in the nano- and picoliter range (4).

5. NUTRIENT ASSAYS

The assays are housed in submicroliter droplets on siliconized microscope slides under heavy mineral oil (heavy white grade, Sigma Chemical Co., St. Louis, MO). For each assay, reagent cocktail solutions are prepared that contain a buffer and all of the co-factors and enzymes needed for the reaction (see Protocols 8.1-8.5). Typically, 10- to 20-nl drops (depending on the substrate to be analyzed) of this reagent cocktail are placed onto a siliconized slide under the mineral oil. The fluorescence of each droplet is measured in turn using a 20x objective by exposing the pyridine nucleotides in the cocktail to the UV light source. Drops are routinely exposed for up to 0.25 s because there is no detectable photo-oxidation of NADH or NADPH during this time.

Following this initial determination of fluorescence, a 1- to 5-nl sample (depending on the substrate to be analyzed) is added to the reagent cocktail drop. The addition of

Figure 8.2. Micrograph of a 1nl microconstriction pipette.

the substrate initiates the reaction. The drops on the slide are then left until the reaction has gone to completion. This time can vary between 3 min and 1 h depending on the substrate to be analyzed. (This must be determined for each assay for a substrate by taking readings over time to determine when the reaction has reached completion.) The fluorescence of the drops is again determined. The change in fluorescence between the reagent cocktail drop before and after addition of the sample should be linear within the concentrations to be assessed. This is determined with a set of standards run on each day of experiment. Unlike spectrophotometric assays, the Beer-Lambert law does not hold for fluorescence microscopy. Therefore it is necessary to run a new set of standards with each day of experiment and to redetermine that the fluorescence range is linear within the concentration of the substrate. An acceptable linear regression value is typically R > .99 (Figure 8.3). Once a linear standard curve has been achieved, the concentration of substrates from samples can be calculated from this curve. The reactions and assay conditions for assays for pyruvate, lactate, glucose, ammonium, and glutamine are shown in Protocols 8.1-8.5.

6. MEASUREMENT OF NUTRIENT UPTAKES BY INDIVIDUAL EMBRYOS

Embryos are incubated in a medium containing the substrates for measurement at levels typically between 0.5 and 1.0 mM. This medium can either be bicarbonate buffered, in which case the incubation must occur in the CO2 incubator, or HEPES/MOPS buffered, in which case the incubation can occur on a calibrated heating stage. Embryos are typically incubated in this medium for 1-4 h depending on the volume of the drop and the stage of embryo to be analyzed. For all uptake experiments, linear rates of uptakes should be determined whenever possible. This measurement will ensure that there are not alterations in substrate uptake or metabolite release as a result of the culture conditions. It has been demonstrated that the culture environment can significantly alter glucose metabolism in just 3 h (5).

For incubation of the embryos, drops of metabolic incubation medium are placed under oil in a dish. Use dishes with low sides, or use the lid of a 35-mm Petri dish. For mouse embryos it is typical to incubate embryos in drops of medium in the 20-50 nl range, while for domestic animal embryos or human embryos volumes of 200 nl to 1 ml are used. Wash embryos well in the metabolic incubation medium to ensure there is no carryover of substrates to the small incubation drops. The embryos are then picked up in a very small amount of volume using a pulled pipette just larger than the embryos.

Lactate Dehydrogenase Pyruvate + NADH + H+ -► Lactate + NAD

Lactate Dehydrogenase Pyruvate + NADH + H+ -► Lactate + NAD

Pyruvate Concentration (mM)

Figure 8.3. Standard curve for pyruvate. Levels of pyruvate in a sample can be assessed by a linear decrease in fluorescence with an increase in pyruvate concentration.

This will ensure that there is a minimal carryover of extra volume into the drops (6). The amount of volume that is carried over into the drops can be determined using triti-ated water.

For determination of linear rates of uptakes or production, serial samples are taken at 20-45 min intervals. A minimum of three readings is required to determine linear rates. For endpoint determinations, an extra three or four drops of medium alone should be included in the incubation to determine the exact amounts of nutrients that were present in the medium and to control for any breakdown that may have occurred during the incubation period. Ensure that the concentration of substrate to be measured does not fall below the Km of the transport system for the substrate. If the samples are not to be analyzed immediately, the embryos can be removed from the drops at the end of the incubation and the media taken up in 1-5 ml capillary tubes surrounded by oil on each end to avoid evaporation. These can then be stored in plastic insemination straws and stored at -80°C. There is negligible breakdown of most substrates during storage for several days under these conditions.

7. ENZYME ANALYSIS

Energy metabolism is controlled by precise regulation of a large number of enzymes. Cells can control the function of enzymes by regulating the availability of cofactors and substrates and by feedback inhibition. Such dynamic changes in metabolic regulation rarely occur by regulating mRNA expression levels or by protein synthesis. Cells can also regulate enzyme activity by changing the isoform of an enzyme. As different isoforms have different kinetic properties, changes in the abundance of a particular isoform can also regulate the flux of a substrate through an enzyme and therefore control pathway activity. Therefore the analysis of enzymes and their kinetics is fundamental to understanding how cells regulate their metabolism and development. Compared to somatic cells, our knowledge of the regulation of enzymes in embryos and how they control metabolism is quite limited. Most of the studies that have been performed have only established the presence of the enzymes and their maximal activity in the presence of nonlimiting conditions (zero-order kinetics). Most often this analysis has been a single reading at the end of an extended incubation and has not used continuous kinetic analysis. The reason for this is the small amount of material present in embryos, which has necessitated the use of complicated procedures such as enzyme cycling to detect enzyme activity (7-9). However, in more recent years, with the further development of microfluorimetry procedures and the increased sensitivity of fluorescence equipment, particularly those with optical switches, it is now possible to establish the kinetics (Km, Vmax) of both cytoplasmic and mitochondrial enzymes in single cells. The procedures described below outline how to determine the kinetic activity of a single enzyme from either a single cell or embryo.

8. ENZYME KINETICS

Enzyme kinetics can be complicated by the various control mechanisms in a cell such as feedback inhibition. The measurement of enzyme activity and establishment of kinetic data can be simplified by making the measurement conditions such that these control mechanisms are eliminated in the measurement conditions. For the assays and procedures listed below, continuous kinetic analyses are used in zero-order kinetics to establish enzyme activity. Using this type of analysis, a single factor (such as the concentration of the substrate or the level of an effector) can be altered in the assay conditions, and the direct effect on enzyme activity can be determined.

Two parameters, Km and Vmax, describe the kinetic properties of an enzyme. The Km of an enzyme is the concentration of the substrate that enables the enzyme to function at halfmaximal activity and is therefore a measure of the specificity of a substrate for the enzyme. For example, the Km of lactate dehydrogenase for pyruvate in mouse embryos is 0.23 mM (10). The lower the Km for a substrate, the higher the specificity of the substrate for the enzyme. Different isoforms of an enzyme have differing values of Km for a given substrate. Therefore, a change in Km can indicate a change in the relative abundance of different isoforms that can alter pathway activity. For example, the glycolytic enzyme hexokinase IV (glucokinase), which is the isoform predominantly found in the liver, has a Km about 250 times higher (~ 10 mM) than the isoforms I-III. Isoform IV is able to control metabolism in the presence of very high concentrations of glucose that may arise in the liver.

The Km for hexokinase in mouse embryos changes from the zygote to the blastocyst stage, indicating a change in the distribution of different isoforms in the embryo as development proceeds (11). This change in enzyme isoform may enable the embryo to alter the activity of the glycolytic pathway. The second parameter that characterizes the kinetics of an enzyme is Vmax or maximal velocity. Both Km and Vmax are determined by establishing the activity of the enzyme at different substrate concentrations using continuous rate measurements (see below). The significance of determining enzyme kinetics is that Km and Vmax can establish the specificity of an enzyme for its substrates and determine the type of reaction that the enzyme controls between steady-state and equilibrium control. This information thereby enables one to determine the contribution of the particular enzyme to the control of a metabolic pathway.

Continuous rate measurement and establishment of kinetics has rarely been performed for enzymes in the mammalian embryo. Instead, velocity has been reported at a single concentration of substrate after several minutes (9, 12-19). Although this measurement ascertains that there is some activity of the enzyme, it tells little about the actual activity of the enzyme and tells nothing about its regulation or importance in the regulation of overall pathway activity. For most enzymes in the mammalian embryo, this highly valuable information is yet to be collected. Measurement of the actual kinetics of an enzyme will provide information about the action of an enzyme within a pathway. Further, it can establish how the enzyme's activity changes in response to changes in substrate availability and the presence or absence of specific effectors (10). This information is essential for understanding how the embryo controls the changes in metabolic activity from a carboxylic acid-based metabolism at the zygote stage to a glucose-based metabolism at the blastocyst stage throughout development.

9. ANALYSIS OF ENZYME ACTIVITY IN EMBRYOS

Activity and kinetics of enzymes in individual cells/embryos can be assessed using ultramicrofluorometric analysis. Similar to the nutrient assays described above, the analysis of enzymes are based on the consumption or production of the pyridine nucle-otides NAD(P)H/NAD(P)+ in coupled reactions. The equipment required for the measurement of enzyme activity is similar to that listed in section 3. However, for enzymes it is desirable for the equipment to have an optical switch that can control the levels of exposure of the samples to UV light to nanoseconds and also therefore able to measure real time enzyme activity.

This type of analysis has been used to determine the kinetics of lactate dehydroge-nase (LDH) in the preimplantation mouse embryo (10) and will be used as an example in this chapter. However, numerous enzymes can be measured in the same fashion by coupling the reactions to the pyridine nucleotides. Fluorescent assays for many enzymes can be found in Bergmeyer and Gawehn (1). Similar to the assays for assessing substrate concentrations, these reactions can then be scaled down to measure the activity in single cells using specially constructed micropipettes.

9.1 Extraction of Enzymes

Enzymes are extracted from individual or groups of embryos using the same procedure. The enzymes are extracted by freezing and thawing the embryos in an extraction buffer (19). A detailed description of the extraction procedure used in our laboratory is shown in Protocol 8.6. This extraction procedure, coupled with the ultra-miroflurometric assay, results in dilution of the cell lysate, which makes endogenous regulators ineffectual. Thus the enzyme/factor of interest can be measured without the need to do extensive purifications that would require a large amount of material.

9.2 Analysis of Enzyme Activity

For each enzyme to be analyzed, a reagent cocktail is prepared in a buffer with the optimal pH and concentrations of substrates and cofactors to ensure zero-state kinetics (i.e., all factors are in excess compared to activity of the enzyme to be measured). The composition and preparation of the reagent cocktail for LDH is described in Protocol 8.7. In addition, a reagent cocktail without substrate is prepared as a control to establish baseline autofluroscence and changes in fluorescence.

For kinetic analysis, the reagent cocktail is prepared with four to six different substrate concentrations (see Protocol 8.7). In addition, a standard must be prepared for each day of experimentation to calibrate and calculate the consumption or production of the pyridine nucleotides.

For all enzymes to be measured, the activity of the assay should be validated by determining the activity of an enzyme standard. Standards can be purchased from several suppliers with actual activities of the lot of enzyme provided (e.g., Roche, Sigma). Using the procedure outlined below, a dilution of the purified enzyme (for LDH 1:2000 dilution) can be substituted for the embryo sample and activity measured. The specific activity can then be calculated and compared with the activity provided by the supplier to validate the assay.

9.3 Measurement of LDH Activity

The Km and Vmax for pyruvate of LDH is measured using the following reaction. LDH activity can be followed by a decrease in fluorescence over time as the fluorescent NADH is oxidized to NAD+.

Enzyme extracts are thawed and expelled onto a siliconized glass slide under mineral oil. The extract should be kept on ice until all assays are completed. The assays are housed in droplets under mineral oil on siliconized glass slides. All enzyme activity should be measured at 37°C. This reaction drop is incubated at 37°C for at least 2 min before the initiation of the reaction by the addition of preequilibrated enzyme (sample) or substrate. Upon initiation of the reaction, the change in fluorescence is monitored for several minutes until the reaction has reached a plateau.

A standard curve of NAD(P)H is run before enzyme samples to enable calibration. For LDH a standard curve of NADH concentrations of between 0 and 0.1 mM is used. Standard curves should have a minimum of four concentrations to determine the linear regression with the fluorescence obtained. A linear regression value of R > .995 is advisable to ensure that the calibration will result in accurate enzyme activities.

The change in fluorescence can then be calculated in units of pyridine nucleotide from the standard curve, taking into account the rate of any autofluorescence that may have resulted from the exposure to UV light. This rate of NAD(P)H consumption or production is then plotted against time.

Figure 8.4. Calculation of activity of lactate dehydrogenase. Rates of NADH consumption are determined over a range of pyruvate concentrations, 0.6 mM, 0.3 mM, 0.15 mM, and 0.075 mM. The velocity of lactate dehydrogenase activity is determined by plotting a tangent to the initial velocity and is measured as picomoles NADH consumed/hour.

Figure 8.4. Calculation of activity of lactate dehydrogenase. Rates of NADH consumption are determined over a range of pyruvate concentrations, 0.6 mM, 0.3 mM, 0.15 mM, and 0.075 mM. The velocity of lactate dehydrogenase activity is determined by plotting a tangent to the initial velocity and is measured as picomoles NADH consumed/hour.

To assess the kinetics of the enzyme, this is repeated on each sample at different substrate concentrations. For determining the Km of an enzyme, use 4-6 concentrations to obtain accurate calculations.

From each embryo sample, it is possible to obtain six different enzyme activity curves for the six different concentrations used (Figure 8.4). These data can then be used to determine a Vi for each concentration. This is the initial velocityV) of the enzyme and can be determined by drawing a tangent to the initial linear portion of the curve. It can be seen from the example in Figure 8.4 that this type of analysis yields vastly different results from endpoint calculations taken after several minutes. Using this approach a V; can be obtained for each of the substrate concentrations used. This can then be plotted as V; against substrate concentration. This graph will produce a Michaelis-Menton plot where the velocity of the reaction is dependent on the substrate concentration.

This confirms that the zero-order kinetics was correct and that there was no carryover of endogenous regulators. However, it is not possible to determine Km or Vmax from these plots. Instead, to determine the kinetic parameters of the enzyme, the equations must be transformed to give either a Lineweaver-Burk or Eadie-Hofstee equations that result in convenient graphical representation of the data that enables easy calculation of Vmax and Km.

The Lineweaver-Burk plot is the most straightforward of these equations. This results in a double-reciprocal graph that plots 1/V against 1/[S]. The slope of this graph is Km/Vmax, and the x-intercept gives -1/Km and the y-intercept gives 1/Vmax (see Figure 8.5). This procedure relies on few assumptions and therefore is beneficial over the least square means analysis, which makes assumptions that are difficult to verify with the small amount of enzyme available from a single embryo.

Using this analysis it is possible to determine a Km and Vmax for an enzyme for each individual embryo. The parameters of Km and Vmax are important because they provide information such as

Figure 8.5. Lineweaver-Burk plot for determination of Km. Rates of lactate dehydrogenase activity at increasing levels of pyruvate are plotted on a double reciprocal plot. The linear regression of this line is r = .998. The equation of the line is y = 0.049x + 0.53. The y-intercept of graph the 1/Vmax, and the x-intercept is -1/Km.

Figure 8.5. Lineweaver-Burk plot for determination of Km. Rates of lactate dehydrogenase activity at increasing levels of pyruvate are plotted on a double reciprocal plot. The linear regression of this line is r = .998. The equation of the line is y = 0.049x + 0.53. The y-intercept of graph the 1/Vmax, and the x-intercept is -1/Km.

1. Determining if there are changes in enzyme isoforms

2. Characterizing the specificity of an enzyme for a substrate (ratio of Vmax/Km is highest for the substrate with the highest specificity)

3. Deciding between steady-state and equilibrium mechanisms

4. Assessing the type of inhibition by negative effectors

5. Indicating the role of an enzyme in metabolism (by relating the Km to the concentration of substrate present in the cell, it is possible to determine the relative activity of the enzyme present).

10. DETERMINATION OF ENZYME ISOFORMS

Many enzymes have different forms, or isoforms. Isoforms arise from genetically determined differences in amino acid sequences. Different isoforms of the enzyme have different kinetic properties, and therefore a change in the isoform present can change the flux through an enzyme and therefore pathway activity. Therefore, the determination of which isoform of an enzyme that is present in a cell is essential for the interpreting and understanding the control of pathway activity. For embryos, there is little information about the isoform of an enzyme that is present throughout development. The most notable exception is for LDH, which was first shown to change from isoform I (heart type) during the preimplantation period in the mouse to isoform V (muscle form) during the peri-implantation period more than 30 years ago (20, 21).

Different isoforms of an enzyme can be separated by electrophoresis. Isoforms are named based on the extent of migration in an electric field, starting with the species with the greatest mobility toward the anode. In this manner it is possible to distinguish between isoforms in different tissues/embryos by examining their migration by electrophoresis.

For embryos, we have found that the best procedure for separation of isoforms is electrophoresis in a 6% acrylamide gel (Protocol 8.8), followed by a staining protocol customized for the individual enzyme using either tetrazolium or diazonium salts. Two extensive books have been published that contain staining protocols for a wide variety of enzymes that can be used for staining isoforms of embryos (22, 23). This method of staining is based on the detection of the activity of the enzymes themselves. Therefore, it is not necessary to purify the sample because only the reaction specific to the enzyme of interest will stain on the gel. This approach enables the detection of isoform patterns of embryos. An example of the staining procedure for LDH is shown in Protocol 8.9. We have successfully used these protocols for staining isoforms for many different enzymes and for distinguishing between cytosolic and mitochondrial enzymes.

Protocol 8.1. Preparation of pyruvate assay reagents and standards

Buffer

2.52 g EPPS 10 mg penicillin 10 mg streptomycin

Make up to 200 ml and adjust pH to 8.0 with 1 M NaOH.

Reagent cocktail (can be frozen in aliquots)

14 ml Buffer

0.4 ml lactate dehydrogenase

Make 5mM NADH by adding 17.7 mg in 5 ml water. Standard curve preparation

Prepare a 1 mM pyruvate solution by dissolving 0.0110 g pyruvate in 100 ml water. Serially dilute the 1 mM pyruvate solution to give final concentrations of 0, 0.0625, 0.125, 0.25, and 0.5 mM. Prepare these solutions daily and use to generate a standard curve.

Protocol 8.2. Preparation of lactate assay reagents and standards

Buffer

7.5 g glycine 5.2 g hydrazine 0.2 g EDTA

Dissolve in 49 ml of water; add 51 ml 2 M NaOH.

Reagent cocktail (prepared day of assay)

0.45 ml Buffer

0.40 ml Water

25 ml lactate dehydrogenase

75 ml NAD+ solution

Make the NAD+ solution 40 mg/ml and store as 75 ml aliquots in freezer. Standard curve preparation

Prepare a 1 mM lactate solution by dissolving 0.0112 g lactate in 100 ml water. Serially dilute the 1 mM lactate solution to give final concentrations of 0, 0.0625, 0.125, 0.25, and 0.5 mM. Prepare these solutions daily and use to generate a standard curve.

Protocol 8.3. Preparation of glucose assay reagents and standards

Hexokinase

Glucose + ATP ^ glucose-6-phosphate + ADP

Glucose-6-phosphate dehydrogenase Glucose-6-phosphate + NADP+ ^ 6-phosphogluconate + NADPH + H+

Buffer

2.52 g EPPS 10 mg Penicillin 10 mg Streptomycin

Make up to 200 ml and adjust pH to 8.0 with 1 M NaOH.

Reagent cocktail (can be frozen in aliquots)

15 ml Buffer 2 ml 5 mM DTT

2 ml 37 mM MgSO4 1 ml 10 mM ATP

3 ml 10 mM NADP

1 ml hexokinase/glucose-6-phosphate dehydrogenase

Compound Dilution

5 mM DTT 7.72 mg in 10 ml water

37 mM MgSO4 91.2 mg in 10 ml water

10 mM ATP 30.3 mg in 5 ml water

Standard curve preparation

Prepare a 1 mM glucose solution by dissolving 0.0180 g glucose in 100 ml water. Serially dilute the 1 mM glucose solution to give final concentrations of 0, 0.0625, 0.125, 0.25, and 0.5 mM. Prepare these solutions daily and use to generate a standard curve.

Protocol 8.4. Preparation of ammonium assay reagents and standards

Glutamate dehydrogenase a-ketoglutarate + NADH + H+ ^ Glutamate + H2O + NAD+

Buffer

9.3 g triethanolamine (TEA)-HCL

95 mg ADP

670 mg a-ketoglutarate

Dissolve in 70 ml water; adjust pH to 8.0 with 5 M NaOH. Make up to 100 ml.

NADH solution

10 mg NADH 20 mg NaHCO3

Dissolve in 2 ml water.

Reagent cocktail (prepared day of assay)

250 ml Buffer 500 ml Water

25 ml glutamate dehydrogenase 25 ml NADH solution

Standard curve preparation

Prepare a 10 mM ammonium chloride solution by dissolving 0.053 g ammonium chloride in 100 ml water. Dilute by adding 1 ml of the 10 mM solution to 9 ml of water. Serially dilute the 1 mM solution to give final concentrations of 0, 0.0625, 0.125, 0.25, and 0.5 mM. Prepare these solutions daily and use to generate a standard curve.

Protocol 8.5. Preparation of glutamine assay reagents and standards

Step 1

Glutaminase Glutamine + H2O ^ Glutamate + NH3

Buffer

6.8 g sodium acetate in 100 ml H2O

2.9 ml acetic acid in 97.1 ml H2O 2.9

Mix 68 ml of sodium acetate with 32 ml of acetic acid. Check that the pH is 5.0.

Reagent cocktail

40 ml Buffer 20 ml Water 20 ml Glutaminase

Dissolve 0.5 mg glutaminase into 1 ml of acetate buffer and freeze in aliquots at -20°C.

Step 2

Glutamate dehydrogenase Glutamate + H2O + NAD+ ^ a-ketoglutarate + NADH + NH4+

Buffer

7.5 g glycine 5.2 g hydrazine 0.2 g EDTA

Dissolve in 49 ml of water, add 51 ml 2 M NaOH.

For 33.5 mM ADP, add 14.3 mg ADP in 1 ml water. For 27 mM NAD+, 18.0 mg in 1 ml water.

Reagent cocktail

200 ml Buffer 20 ml NAD+ 10 ml adp

50 ml glutamate dehydrogenase Standard curve preparation

Prepare a 1 mM glutamine solution by dissolving 0.0147 g glutamine in 100 ml water. Serially dilute the 1 mM glutamine solution to give final concentrations of 0, 0.0625, 0.125, 0.25, and 0.5 mM. Prepare these solutions daily and use to generate a standard curve.

Protocol 8.6. Enzyme extraction from embryos

Extraction buffer

100 mM K2HPO4 30 mM KF

1 mM EDTA

5 mM P-mercaptoethanol

Adjusted to pH 7.5.

Phenylmethylsulfonyl fluoride (PMSF) stock

Dissolve 10 g/l PMSF in ethanol and store at 4°C. Immediately before use, add 50 ml of the PMSF stock to 950 ml of the extraction buffer.

1. Wash individual or groups of embryos in saline supplemented with 4 mg/ml BSA.

2. Transfer embryos in a minimal amount of volume (<1 ml) into a 500 ml drop of the extraction buffer.

3. Pick up embryos in a 200-nl to 1-ml drop of the extraction buffer and place the drop containing the embryos under oil.

4. Using a washed glass capillary (1-5 ml, Drummond) take up the drop containing the embryo surrounded on either end with oil, which will prevent evaporation.

5. Seal the glass capillary within a plastic insemination straw and kept frozen at -80°C until analysis.

Note: For some enzymes such as phosphofructokinase the enzyme is not stable stored at -80°C and therefore should be immediately thawed and analyzed.

Protocol 8.7. Preparation of lactate dehydrogenase reagents and standards

Buffer

K2HPO4 34.0 mM KH2PO4 7.3 mM

For kinetic analysis, prepare 5 solutions containing pyruvate:

1. 100 ml buffer containing 1.25 mM pyruvate

2. 100 ml buffer containing 0.63 mM pyruvate

3. 100 ml buffer containing 0.31 mM pyruvate

4. 100 ml buffer containing 0.16 mM pyruvate

5. 100 ml buffer containing 0.0.8 mM pyruvate

NADH solution

Prepare the 11.3 mM NADH solution fresh daily. Add 9.3 mg NADH and 10 mg NaHCO3 to 1 ml of water and dissolve.

Assay cocktail

For each buffer solution, a cocktail solution is prepared consisting of 1 ml buffer and 5 ml NADH solution.

Standard curve preparation

Prepare a 10 mM NADH solution by dissolving 7.1 mg NADH into 1 ml water. Dilute by adding 10 ml of 10 mM solution to 990 ml of water to give a 100 mM NADH solu tion. Serially dilute the 100 mM NADH solution to give final solutions of 0, 3.125 mM, 6.25 mM, 12.5 mM, 25.0 mM, 50 mM, and 100 mM. Prepare these solutions daily and use to generate a standard curve for calibration of lactate dehydrogenase activity.

Protocol 8.8. Preparation of a 6% acrylamide gel

Component

10 ml

20 ml

30 ml

50 ml

H2O

5.3

10.6

15.9

26.5

30% Acrylamide mix

2.0

4.0

6.0

10.0

1.5M Tris base (pH 8.8)

2.5

5.0

7.5

12.5

10% Ammonium persulfate (w/v)

0.1

0.2

0.3

0.5

TEMED

8 ml

16 ml

24 ml

40 ml

Mix all the solutions except for the TEMED into a glass flask. Add the TEMED immediately before the gel is poured.

Mix all the solutions except for the TEMED into a glass flask. Add the TEMED immediately before the gel is poured.

Running buffer

Dissolve 4 g glycine and 1.2 g Tris base in 2 l water and adjust pH to 8.3. Run gel at 40 amps (around 250 V) for 1.5 h. Keep the gel cool by either running in the cold room or by cooling with cold water.

Protocol 8.9. Staining solution for lactate dehydrogenase isoforms

Staining of the gel relies on the activity of the enzyme. The gel is stained using the dye combinations of tetrazolium salt nitro-BT.

Phenazine methosulfate (PMS) acts by accepting a hydrogen from the NADH and passing it to the tetrazolium salt nitro-BT. When the nitro-BT is reduced, the dye is blue-black, and therefore wherever there is LDH activity it will appear as a blue-black band.

Lactate Pyruvate

Lactate Pyruvate

Formazan Nitro-BT

(blue/black color)

Staining solution

0.2 M Tris-HCl buffer, pH 8.0 0.5 M lactate 10 mg/ml NAD+ 1 mg/ml nitro-BT 2.5 mg/ml PMS

Incubate gel in staining solution for about 1 h in the dark at 37°C or until bands appear.

References

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2. Gardner, D. K., and Leese, H. J. (1999). In: Handbook of in Vitro Fertilization, 2nd ed., A. O. Trounson and D. K. Gardner, eds., p. 347. CRC Press, Boca Raton, FL.

3. Gardner, D. K., and Leese, H. J. (1993). In: Handbook of in Vitro Fertilization, A. Trounson and D. K. Gardner, eds., p. 195. CRC Press, Boca Raton, FL.

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14. Brinster, R. L. (1966). Exp. Cell Res., 43, 131.

15. Brinster, R. L. (1967). Biochim. Biophys. Acta, 148, 298.

17. Brinster, R. L. (1971). Wilhelm Roux Arch. Entwicklungsmech. Org., 166, 300.

18. Brinster, R. L. (1968). J. Reprod. Fertil., 17, 139.

19. Martin, K. L., Hardy, K., Winston, R. M., and Leese, H. J. (1993). J. Reprod. Fertil., 99, 259.

20. Auerbach, S., and Brinster, R. L. (1967). Exp. Cell Res., 46, 89.

21. Auerbach, S., and Brinster, R. L. (1968). Exp. Cell Res., 53, 313.

22. Rothe, G. M. (1994). Electrophoresis of Enzymes. Springer Verlag, Berlin.

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