TY - JOUR
AU - Zhang, Shuqiu
AB - Abstract The unique roles of individual cells may be critical to the physiology of an organism. In such cases, micromethods are essential to elucidating the molecular biology, biochemistry and biophysics of the specialized cells or even subcellular compartments of the important cells. The great proliferation of micromethods testifies to their value and no single review can be comprehensive. This review therefore provides only a generalized overview of one approach, namely dissection that provides a pure sample for subsequent extraction and analysis by microdroplet chemistry. As a means of illustrating the utility of this approach, an application—study of the interaction of cytosolic malate concentration and guard‐cell phosphoenolpyruvate carboxylase—is provided. Cellular localization, compartmentation, guard cells, histochemistry, individual cell, malate, micro, phosphoenolpyruvate carboxylase, single cell, stomata. Introduction Often, tissue heterogeneity confounds an interpretation of results obtained by organ‐level or tissue‐level analyses. For example, guard cells act semi‐autonomously in the leaf surface and control two of the most important physiological operations of the plant, namely, acquisition of CO2 and regulation of water loss. Yet these cells comprise only a tiny fraction of the whole leaf and even a tissue‐level analysis would reveal little about guard cells per se. In other cases, exemplified by C4 photosynthesis, abundant but disparate cells contribute uniquely to a single function. Thus, a leaf‐level analysis would not permit the assignment of one or another task to either mesophyll cells or bundle‐sheath cells. These two examples suffice to illustrate the well‐known requirement to conduct investigations at the cell level in many instances. Many sensitive methods have therefore been developed. In all cases, the first decision is to adopt a strategy for tissue sampling. Strategies for obtaining single‐cell‐size samples fall into one of two general categories. First, in situ methods (e.g. enzyme histochemistry, RNA hybridization) rely on localization of an indicator to a cellular or subcellular region that remains in the context of the tissue. Second, other methods depend on the removal of the sample from the tissue context. A well‐known example of this basic strategy is protoplast isolation, but another method is to remove a cell‐size sample for analysis. The cell‐size sample may be taken directly via a capillary (Sims et al., 1998) or cellular contents may be removed via pipettes (Karrer et al., 1995; Koroleva et al., 1998). Alternatively, the tissue may be stabilized and cells or subcellular samples subsequently dissected out. Regardless, separation of the sample from the tissue before analysis facilitates simultaneous multiple analyses, such as use of separation or array technologies. For example, a large number of mRNA‐abundance patterns in the extract of a single sample can be monitored at once, impossible by the analogous in situ method. Monitoring a large number of analytes simultaneously is not only efficient, but permits assignment of new functions to known molecules (Brent, 2000) or ions. Simple manual dissection of cells or subcellular samples is a simple, low‐technology and inexpensive means of obtaining a small specimen for analysis by microdroplet chemistry. Curiously, this method is rarely used in plant‐cell biology. The purpose of this article is to suggest a wider application of hand dissection, which should be considered as a starting point for single‐cell biochemistry or molecular biology. Application examples are given that emphasize the value of manual dissection. The focus is on sample preparation, handling and extraction, all of which are common to subsequent analyses of various sorts. Tissue preparation Cryofixation Rapid freezing of plant samples is adequate to preserve most metabolites and enzymes. Generally, if liquid nitrogen is cooled from its boiling point to its melting point by application of a vacuum, a small sample held by the edge with thin forceps can be plunged into liquid nitrogen slurry without the formation of insulating air bubbles. Any experimental design depends on the purpose at hand, and general procedures may be inadequate. As an example, ATP concentrations in vivo are affected by oxygen, and ATP pools turn over rapidly (Oresnik and Layzell, 1994). Furthermore, ATP is susceptible to hydrolysis during tissue disruption, freezing, and freeze‐drying (Trautschold et al., 1985) and during storage at −20 °C (Passonneau and Lowry, 1993). In such cases, alternative coolants or methods that provide ultra‐rapid freezing by high‐pressure fluid (Craig and Staehelin, 1988) or pre‐cooled metal tongs should be considered. Stabilization for dissection Freeze‐drying (removal of tissue water by sublimation at low temperature and pressure, see Passonneau and Lowry, 1993) and freeze‐substitution (removal of tissue water by dissolution into an organic solvent at low temperature, see Parthasarathy, 1995) are generally adequate means of preserving the chemical integrity and localization of plant analytes. With these methods, even a cold‐labile, light‐activated enzyme activity, pyruvate, orthophosphate dikinase (EC 2.7.9.1), has been preserved in dissected cells of a C4 plant (Outlaw et al., 1981). Similarly, early studies (Fisher and Outlaw, 1979) showed that the intracellular localization of water‐soluble metabolites such as 14C‐labelled products of photosynthesis could be maintained. However, de Brock's caution that freeze‐drying and freeze‐substitution are ‘cumbersome and often give poor results with plant material’ merits consideration (de Brock, 1995). First, in the case of freeze‐drying, high standards (<−35 °C, ≤10 μm Hg) must be maintained throughout the process. Second, in the case of freeze‐substitution, rigorous and strictly anhydrous procedures must be followed. Even under the best conditions, each procedure must be evaluated for artefacts. For example, a fraction of the activity of sucrose synthase (EC 2.4.1.13) was lost during freeze‐drying (Hite et al., 1993) as judged by comparisons of the activity in fresh tissue extracts. As another example, 6% of photosynthetically formed 14C‐phosphoglycerate was hydrolysed to glycerate during freeze‐substitution and embedment (Outlaw and Fisher, 1975b). Dissection Tissue stabilization is the first goal of any dissection protocol. Tissues that are easy to fragment, such as Vicia faba leaves, can be freeze‐dried intact and subsequently dissected. Compact, cohesive tissues, such as Dianthus meristem, may be cryosectioned before freeze‐drying and dissection (Croxdale and Outlaw, 1983). If tissue is freeze‐substituted and embedded, it must be sectioned before dissection. Regardless of the method, stabilized tissues must be handled in low humidity to avoid artefacts of diffusion or metabolism. Under these controlled conditions after stabilization, no plant metabolite has been reported to be degraded. The most labile plant enzyme documented for these conditions is glucose‐6‐P dehydrogenase (EC 1.1.1.49), which has a half‐time of 30 h (Croxdale and Outlaw, 1983). Thus, although each tissue and each analyte should be evaluated, no loss of practical importance has been ascribed to ambient conditions during dissection. Morphological resolution is the second goal of any dissection protocol. When coherent masses of a single‐cell type, such as palisade parenchyma, are present, relatively large masses can be dissected easily. In other cases, it is possible to tease out the desired structure (representative references in Zhou et al., 2000), but usually, excision is required. Therefore, laser microdissection was developed early (Meier‐Ruge et al., 1976), but the technology did not become widely used, probably because of artefacts associated with heat required to burn the tissue into sections. However, interest in this general technology has mushroomed recently (Bonner et al., 1997; Schlindler, 1998; Simone et al., 1998; Suarez‐Quian et al., 1999) with the development of laser capture microdissection. In essence, laser capture microdissection depends on overlaying a 5 μm thick cryopreserved section with thermoplastic film. An infrared laser is focused on the area of interest (resolution ∼1 μm, Simone et al., 1998), melting the polymer and embedding the tissue. When the area of interest is lifted, the adjacent, unembedded tissue shears away. Thus, laser‐capture microdissection is not dissection in the usual sense of the word, and this technique does not burn through the edges of a tissue section as previous laser‐based methods did. The aggressive marketing of this method has had the positive effect of focusing attention on the importance of cell‐specific physiology, biochemistry and molecular biology, but the positive attributes of simple manual dissection (Passonneau and Lowry, 1993) are vastly understated in the market (http://www.arctur.com/faqs.html). Hand‐dissection can be simply carried out by use of a razor blade fragment mounted onto a handle (Passonneau and Lowry, 1993). Indeed, manual or assisted dissection of specific animal cells has a long history (Lowry, 1973) and finds current application in medicine and molecular biology (Cannizzaro, 1996; Macintosh et al., 1998) and biochemistry (Teutsch et al., 1995). In the specific case of higher plants, cells such as palisade cells can be hand dissected cleanly (Fig. 1), and the general limit of hand dissection is about 2 μm (Outlaw, 1980). As the examples in the following sections show, hand dissection permits the study of intra‐ and intercellular compartmentation with picolitre resolution. It is important to note, however, that many biological studies do not require the resolution illustrated in this article. Fig. 1. Open in new tabDownload slide Nanogram‐size cell samples that were hand dissected from freeze‐dried Vicia faba leaflet. (A) Guard‐cell pair (∼6 ng), dissected as a unit from the abaxial epidermis. (B) Abaxial epidermal‐cell sample. (C) Palisade cell (∼12 ng). (D) Spongy cell (∼14 ng). The scale bar is 20 μm. Dry cell masses are from Jones et al. (Jones et al., 1977) and Outlaw and Lowry (Outlaw and Lowry, 1977). (Reproduced with permission of Springer‐Verlag; from Hampp R, Outlaw Jr WH, 1987. Mikroanalytik in der pflanzlichen Biochemie. Naturwissenschaften74, 431–438.) Basis for analyte expression Overview Interpretation of the analyte content of microdissected samples requires a specific biological basis for expression. The appropriate basis, for example, protein, chlorophyll, dry mass, cell volume, membrane surface area, depends on the biological question. However, the size of dissected samples is intrinsically limiting and direct microassays of the usual bases, for example, protein (Outlaw, 1995), are destructive and laborious. Fortunately, in some applications, determination on a cell basis is adequate. In other applications, a simple non‐destructive method for determining mass or volume, as described below, is required. Subsequently, the cell, mass, or volume expression is converted to the relevant basis. Determination of mass of microdissected freeze‐dried samples by the quartz fibre ‘fish pole’ balance Developed in 1939 by Lowry when he was working with Linderstrom‐Lang, the quartz fibre balance was continually refined over the following 40 years when a safe source of radiation (sealed 241Am) to dispel static electricity was added (Outlaw Jr WH, unpublished results). As described in Fig. 2, the balance is ingeniously simple and inexpensive to construct. It is also utterly sensitive, capable of determining the mass of sub‐ng samples (Kato et al., 1973). The disadvantages are practical. First, although with care they are relatively easy to use, small balances that are used for mass determinations of single plant cells require finesse to construct. Second, each balance has a small useful linear range (say, a 10‐fold range of masses). Third, they are relatively imprecise compared with other balances. Fig. 2. Open in new tabDownload slide The quartz‐fibre ‘fish pole’ balance, used for weighing nanogram‐size tissue samples. A thin, quartz fibre mounted horizontally at one end inside a reversed glass syringe (A) is the basic component of the balance. The syringe protects against air currents and contains a disc of 241Am that dispels static. The tissue sample is transferred to the balance fibre from a sample holder (B), which is visualized through an horizontally mounted dissecting microscope (C). The mass‐induced deflection of the balance fibre toward gravity (D) is measured with the same microscope (E). The balance housing and the microscope are mounted onto the same vibration‐dampened slab of stone. (Revised and reproduced with permission of Humana Press; from Passonneau JV, Lowry OH, 1993. Enzymatic analysis. A practical guide. Totowa, Humana Press.) Determination of volume of freeze‐substituted, embedded microdissected samples by fluorescence Outlaw and Fisher devised a fluorometric assay for the volume of microdissected samples of methacrylate‐embedded tissue (Outlaw and Fisher, 1975a). In brief, the basis of the assay is inclusion of the fluor BBOT ((2,5‐bis‐tert‐butylbenzoxazolyl)‐thiophene) in the methacrylate monomers. Following polymerization, the BBOT is uniformly distributed throughout the interior of the methacrylate block. The sample is sectioned and microdissected; then, the fluorescence of the sample is assayed (excitation: 338 nm; emission: 475 nm). The fluorescence of the sample is compared with the fluorescences of larger reference samples of the same methacrylate block. The masses of the reference samples (quartz‐fibre balance, above) and the density of methacrylate permit the conversion of sample fluorescence to sample volume. Extraction Analysis of microdissected samples requires the use of small volumes in order to diminish the effect of contamination in reagents (‘blank’) and to increase the concentration of the analyte. However, the use of small volumes introduces the problem of reagent evaporation as well as some assay‐specific problems associated with reagent surface area. Fortunately, in protocols requiring >20 nl, evaporation is eliminated by working under oil. It is also possible with extra precautions to control evaporation in much smaller droplets by working under oil or in a humid chamber. Figure 3 describes the oil‐well technique (Passonneau and Lowry, 1993) the basic component of which is a small slender pipette used to deliver reagents into oil‐filled holes. Experience with plant applications (down to 1.5 nl, Outlaw and Kennedy, 1978) has been with hand‐fabricated quartz pipettes (Passonneau and Lowry, 1993), but several automated pipette‐construction protocols (Quinton, 1976) have been published. Overall, however, manipulation of nanolitre droplets is tedious and labour‐intensive and should be replaced by automated systems. Microfluidics (aliquoting, transporting and merging microdroplets) is a big challenge, but biology should be able to draw from other sciences (e.g. piezoelectric ‘ink‐jet’ systems for the delivery of small volumes). Current work (Washizu, 1998; Cooper, 1999; Jones TB, personal communication) demonstrates both progress and difficulties in the development of the appropriate technologies. As an example, Jones (TB Jones, personal communications) has been able to produce and merge 7 nl droplets using electrostatic forces, which are adaptable to microprocessor automation. When perfected, this level of miniaturization should be adequate for most purposes. Fig. 3. Open in new tabDownload slide The ‘oil‐well’ technique, used for analysing nanogram‐size tissue samples. A series of cylindrical 3 mm holes, drilled into a 5 mm thick Teflon block, are partially filled with oil. Extraction and some subsequent assay steps are conducted under oil (see text), which eliminates or reduce evaporation of the reagent. A slender, pointed, constriction pipette delivers an aliquot of extracting solution (A). A tissue sample is pushed through the oil into contact with the extracting solution (B). Subsequently (C), aliquots of other solutions may be added or removed from the oil well, as appropriate for the particular assay. The volumes shown are for perspective and depend on assay design. (Revised and reproduced with permission of Humana Press; from Passonneau JV, Lowry OH. 1993. Enzymatic analysis. A practical guide. Totowa, Humana Press.) Example applications Overview: technical approaches Two complementary applications will illustrate use of hand dissection followed by microdroplet chemistry. The first uses freeze‐drying (for a whole‐cell sample) and the second, freeze‐substitution (for a subcellular sample). Fortuitously, both rely on measurement of oxidation of NADH either by real‐time microdroplet fluorometry (first example, an enzyme assay) or by indirect end‐point analysis by enzymatic cycling (second example, a metabolite assay). As NAD(P) can be coupled directly or indirectly to most enzymatic reactions, generic methods of measuring the oxidation of NAD(P)H or reduction of NAD(P)+ can be adapted for varied purposes. Overview: biochemical context of analyses Post‐translational regulation by reversible protein phosphorylation of phosphoenolpyruvate carboxylase (PEPC, EC 4.1.1.31) isoforms in C4 and CAM cells has long been known (Chollet et al., 1996; Vidal and Chollet, 1997; Nimmo, 2000). This regulatory phosphorylation is manifested kinetically as reduced sensitivity to malate inhibition under suboptimum assay conditions that are presumed to mimic the physiological milieu. PEPC also plays a central role in stomatal movements (Outlaw, 1990; Asai et al., 2000), and guard cells contain a specific isoform (Schulz et al., 1992; Wang et al., 1994; Nast and Müller‐Röber, 1996). In contrast to the photosynthetic isoforms, specific regulatory phosphorylation of guard‐cell PEPC was not detected in guard‐cell protoplasts (Schnabl et al., 1992). Moreover, a direct measurement of cytosolic malate concentration in undisturbed plant tissue was lacking (but see Steingraber and Hampp, 1987; Chang and Roberts, 1989). Thus, the example analyses below focus, first, on in vitro inhibition of guard‐cell PEPC by malate and, second, on malate concentration in plant cytosol. Information obtained from both investigations is necessary to an understanding of how the accumulation of malate in guard cells during stomatal opening is regulated. Activation state of guard‐cell phosphoenolpyruvate carboxylase in relation to the physiological status of the leaf This section describes a study of enzyme kinetics in dissected cells using NADH fluorescence in microdroplets as the reaction indicator. Other uses of microfluorometry having cellular and subcellular resolution that do not rely on hand dissection will not be discussed. Valuable and widespread, these other methods include quantitative in situ autofluorescence kinetics analysis of chlorophyll (Vaughn and Outlaw, 1983; Oxborough and Baker, 1997; Baker et al., 2001) and pyridine nucleotides (Griffiths et al., 1998) as well as analysis of ion‐sensitive fluorescence of xenobiotics (McAinsh and Hetherington, 1998). It is also noted that many artificial fluorogenic substrates have been developed that provide alternative methods and extend the scope of enzymatic reactions that can be studied quantitatively with small dissected samples (Haugland, 1995; Gee et al., 1999). Fluorescence is a more specific means of measuring reduced NAD(P) than absorbance is. In addition, fluorescence is a measurement of absolute light and is thus inherently more sensitive than absorbance, which is calculated from diminution of transmitted light. Finally, the fluorescence signal can be increased within limits because fluorescence is proportional to excitation. Although there are disadvantages to measuring NAD(P)H fluorometrically (e.g. temperature dependence), it is an attractive method when sensitivity is an important aspect of analytical design. Thus, fluorometric methods for measuring pyridine nucleotides and other substances in microdroplets were developed (Rutili et al., 1976; Mroz and Lechene, 1980; de Josselin de Jong et al., 1980, and references therein) as instrumentation became available. Building on these earlier methods, Outlaw et al. increased the sensitivity ∼100× through optimization of the optical system and dedicated software (Outlaw et al., 1985a, b), making it possible to measure single‐cell enzyme activities in real time using natural substrates in solution. Although the analysis described in the following paragraph was produced with custom‐fabricated equipment, currently available turnkey systems (Deutsch et al., 2000) would appear to be easily adapted to freeze‐dried dissected plant cells. The specific application example concerns the kinetics state of guard‐cell PEPC. Aware that enzyme activities in guard‐cell protoplasts can be labile (Hite and Outlaw, 1993) and that the phosphorylation domain of PEPC is easily proteolysed (Chollet et al., 1996), Zhang et al. designed a PEPC assay based on microdroplet fluorometry suitable for analysis of single guard‐cell pairs dissected from freeze‐dried leaf tissue (Zhang et al., 1994). The principle of the assay is real‐time measurement of the PEPC product, OAA, by coupling to malate dehydrogenase. Thus, PEPC activity is indicated by a decline in NADH fluorescence following the addition of the tissue sample to assay cocktail. This assay is chosen for illustration because it is more difficult to perform than the ordinary single‐cell assay (contrast with Tarczynski and Outlaw, 1990). The difficulty stems from the requirement for low NADH concentration (because the assay is conducted under suboptimum conditions) and the need to consume much of the NADH over a short time‐course (to prevent a protein dephosphorylation artefact). An additional complication was introduced by the need for high malate concentration in the presence of endogenous and analytical malate dehydrogenase (cf. Outlaw and Manchester, 1980). Despite these assay constraints, it was easily demonstrated that PEPC in guard cells of opening stomata was insensitive to malate, whereas this negative allosteric effector inhibited PEPC in guard cells of closed stomata (Fig. 4). This alteration in kinetics, which corresponded to a physiological state of the tissue, led to the demonstration that guard‐cell PEPC is reversibly phosphorylated when stomata are stimulated to open (Du et al., 1997; Cotelle et al., 1999). Fig. 4. Open in new tabDownload slide Phosphoenolpyruvate carboxylase (PEPC) activity in individually dissected guard‐cell pairs of Vicia faba (see Fig. 1). PEPC was assayed fluorometrically in real time (Outlaw et al., 1985a) under suboptimum conditions of pH and limiting substrate in a 17 nl droplet. Each trace is an average of seven individual time‐courses; for a single‐assay trace under optimum conditions (see Tarczynski and Outlaw, 1990). PEPC in guard cells of closed stomata (left panel) is inhibited by 400 μM malate whereas PEPC in guard cells of opening stomata (right panel) is insensitive to 400 μM malate. These physiological states correspond to the non‐phosphorylated and phosphorylated forms of guard‐cell PEPC, respectively (Du et al., 1997). (Reproduced with permission of Elsevier; from Zhang SQ, Outlaw Jr WH, Chollet R, 1994. Lessened malate inhibition of guard‐cell phosphoenolpyruvate carboxylase velocity during stomatal opening. FEBS Letters352, 45–48.) Quantification of cytoplasmic malate concentration This section describes a study of subcellular metabolite concentrations in dissected cells using enzymatic cycling. Enzymatic cycling as a means of providing chemical amplification can be traced to Warburg in the 1930s (Lowry, 1990), but it was through OH Lowry's knack and persistence that laboratory protocols became routine. By 1980 (Lowry, 1980), 19 different cycling protocols had been published for NAD+/NADH, NADP+/NADPH, ATP/ADP, GTP/GDP, GSSG/2GSH, and Co A/Acetyl Co A. Currently, various new cycling procedures for pyridine nucleotides (Obon et al., 1999) and other substances (Sakakibara et al., 1999) are being developed, attesting to the currency and importance of this approach. The cycling systems based on NAD and NADP are the most important for two reasons. First, NAD(P) redox reactions, as mentioned, can be coupled directly or indirectly to many enzymes and metabolites. Second, the oxidized and reduced forms of pyridine nucleotides can be selectively destroyed. (For other general information about enzymatic cycling, consult Lowry, 1973; Outlaw, 1980; Passonneau and Lowry, 1993.) The principle of enzymatic cycling is simple (Fig. 5), as further explained as part of the following generalized procedure for a metabolite: In a first or extraction step, the tissue is pushed onto a microdroplet (Fig. 3A, B), which is then heated to destroy endogenous enzymes (and co‐factors in some cases). In a second or specific step, the metabolite is coupled enzymatically to a co‐factor by addition of a second microdroplet (Fig. 3C). The simplest case broadly outlined here involves an analytical dehydrogenase that is specific for the metabolite of interest and the cofactor is NADH. After this step, NAD+ is equal to the amount of metabolite originally present in the extract. In a third or destruction step, analytical enzyme(s) and unreacted co‐factor remaining from the specific step are destroyed. Thus, this example would call for addition of acid, which destroys NADH (but not NAD+). After this third step, the total NAD present is the NAD+ formed stoichiometrically with the reduction of the metabolite in the specific step. In a fourth or cycling step (Fig. 5), the co‐factor is amplified. This and the following reactions are generic—they provide a means of measuring total co‐factor, in this case, NAD. (The principles of cycling are the same for other co‐factors, such as NADP or ATP.) As explained (Fig. 5), NAD+ initiates a cyclic reaction, resulting in the accumulation of products B and D. As the co‐factor is added in low concentration, well below the Km, product accumulation is linear with the amount of co‐factor added.  In a fifth or indicator step, the accumulated product (B or D, Fig. 5) is assayed by a conventional procedure in which the assay volume is in the millilitre range. The specific application example concerns the cytosolic concentration of malate, a negative effector of PEPC. Aware that plant‐tissue malate concentration is heterogeneous, subject to environmental conditions (Gerhardt and Heldt, 1984), and as high as the millimolar range (Steingraber and Hampp, 1987; Chang and Roberts, 1989), Bodson et al. designed a malate assay suitable for use on undisturbed subcellular tissue samples in which PEPC activity and whole‐tissue malate accumulation could be co‐ordinately determined (Bodson et al., 1991). The purpose was to establish whether malate concentration is sufficient to inhibit PEPC as predicted from in vitro kinetics and, if so, whether the in vivo rate is indeed inhibited. The model was Raphanus sativus root hairs (Fig. 6), which were freeze‐substituted. The dense cytoplasmic ‘cap’ permitted the dissection of picolitre volumes of cytoplasm, which was analysed for malate content (Fig. 7). The conclusion was quite clear: high cytosolic malate concentration did not inhibit PEPC in vivo, indicating that PEPC existed in an altered kinetic state in vivo. Although this model was not pursued further, enzyme phosphorylation (see previous section) and presence of allosteric effectors (particularly glc 6‐P) would be the current interpretation of the altered state. The malate analysis above demonstrates the potential utility of dissection and microdroplet chemistry to study metabolite compartmentation between the cytoplasm and the vacuole. In other cases, dissection has permitted the measurement of cell‐wall substances, such as mannitol (by radioactive assay, Ewert et al., 2000), sucrose (by enzymatic cycling, Lu et al., 1997) and ABA (by ultrasensitive immunoassay, Zhang and Outlaw, 2001). Fig. 5. Open in new tabDownload slide The principle of enzymatic cycling. The molecule to be amplified, X, is added to reagent containing excess A and an enzyme that couples the conversion of A to B with the conversion of X to Y. In the same reagent, Y is reconverted to X, which, again, is converted to Y. For each turn of the cycle, B and D accumulate. After hundreds or thousands of cycles, B and D accumulate sufficiently to be measured by conventional means. NAD+/NADH and NADP+/NADPH cycles are the most popular because these co‐factors can be coupled stoichiometrically in a preceding step to a variety of analytes or enzymes. Many other cycling protocols (e.g. for ATP/ADP) are also available and extends the utility of the approach. Fig. 6. Open in new tabDownload slide Emerging root hair of Raphanus sativus. The tissue was quickly frozen, freeze‐substituted at −80 °C, embedded in anhydrous methacrylate, and sectioned. (A) Light micrograph, demonstrating a dense cytoplasmic cap and a contrasting clear vacuole. Such clear visualization tests the limits of hand dissection of distinct regions of a cell. The scale bar is 10 μm. (B) Electron micrograph of the cytoplasmic cap. The scale bar is 1 μm. (Reproduced with permission of the Histochemical Society; from Bodson MJ, Outlaw Jr WH, Silvers SH, 1991. Malate content of picoliter samples of Raphanus sativus cytoplasm. Journal of Histochemistry and Cytochemistry39, 435–440.) Fig. 7. Open in new tabDownload slide The malate content eluted from samples that were hand dissected out of 6 μm sections of methacrylate‐embedded R. sativus root hairs (Fig. 6). The x‐axis is the sample volume determined from the fluorescence of a fluor (BBOT, see text) that was incorporated into the embedding matrix. The closed symbols are from samples of cytoplasmic cap whereas the open symbols are from heterogeneous samples that contained vacuole (61%) and cytoplasm (39%). Results of two experiments, indicated by the different symbols, are shown. The concentration of malate in the cytoplasmic cap was 8.4 mM (or as much as 10 mM, if the volume of the Golgi‐derived vesicles is excluded). The concentration of malate in the vacuole was 54 mM. (Reproduced with permission of the Histochemical Society; from Bodson MJ, Outlaw Jr WH, Silvers SH, 1991. Malate content of picoliter samples of Raphanus sativus cytoplasm. Journal of Histochemistry and Cytochemistry39, 435–440.) Flexibility and accessibility of microanalysis The preceding emphasis on sensitivity limits and morphological resolution dampens equally important messages of this article, assay flexibility and accessibility, so a brief postscript to consider adaptability to a typical laboratory situation is in order. Consider an analysis for the sucrose content of a single palisade cell (Fig. 1), which contains approximately 2 pmol sucrose (Jones et al., 1977): (1) The initial extracting cocktail, 2 μl, is delivered into the bottom of a silanized 6 mm borosilicate tube with a standard commercial repeating pipette, which can deliver volumes as small as 0.1 μl. (2) A palisade cell is pushed onto the 2 μl droplet by means of a small quartz fibre mounted onto a handle. (3) The droplet is covered by oil (40% hexadecane+60% USP light mineral oil) and heated to destroy endogenous enzymes and pre‐existing interfering substances. (4) Additional steps of the assay (to couple sucrose hydrolysis to reduction of NADP+) are conducted by addition of reagents through the oil onto the droplet. (5) An aliquot, 5 μl, is removed for amplification and final measurement in 1 ml. As this outline shows, a microassay can be very simple. Indeed, the only specialized facility for the outlined microassay is an environment suitable for dissecting. Conclusion Plants are successful because biological processes are compartmented at the cellular and subcellular levels. In many instances, these processes can only be understood by investigation of specific compartments. A proven, powerful means of isolating the compartment for biochemical and physiological studies is by dissection followed by microdroplet chemistry. This sampling approach should lend itself equally well to single‐cell gene expression analysis by differential display (Renner et al., 1998) or DNA microarrays (Freeman et al., 1999) and to microcolumn separation methods (Kennedy et al., 1989; Valaskovic et al., 1996; Bächmann et al., 1998; WH Outlaw Jr, H Lochmann, K Bächmann, unpublished results). 1 To whom correspondence should be addressed. Fax: +1 850 644 0481. E‐mail: outlaw@bio.fsu.edu 2 Present address: College of Biological Sciences, China Agricultural University, Beijing, China 100094. The US Department of Energy supported the work in the laboratory during the preparation of this review. References Asai N, Nakajima N, Tamaoki M, Kamada H, Kondo N. 2000 . Role of malate synthesis mediated by phosphoenolpyruvate carboxylase in guard cells in the regulation of stomatal movement. Plant and Cell Physiology 41, 10 –15. Google Scholar Crossref Search ADS PubMed Bächmann K, Lochmann H, Bazzanella A. 1998 . 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TI - Single‐cell dissection and microdroplet chemistry
JF - Journal of Experimental Botany
DO - 10.1093/jexbot/52.356.605
DA - 2001-04-01
UR - https://www.deepdyve.com/lp/oxford-university-press/single-cell-dissection-and-microdroplet-chemistry-eQ027mlx09
SP - 605
EP - 614
VL - 52
IS - 356
DP - DeepDyve
ER -