Biosensors & Bioelectronics 7 (1992) 255-272

Review article Biosensors based on the energy metabolism of living cells: The physical chemistry and cell biology of extracellular acidification John C. Owicki Molecular

Devices Corporation,

& J. Wallace

4700 Bohannon

Parce

Drive, Menlo Park, CA 94025, USA

(Received 2 May 1991; revised version received 14 August 1991; accepted 28 August 1991)

Abstract: The silicon microphysiometer is a biosensor-based instrument that detects changes in the physiological state of cultured living cells by monitoring the rate at which the cells excrete acidic products of metabolism. This paper discusses the chemical and biological factors that determine the performance and applications of such a system. Under typical culture conditions, extracellular acidification is dominated by the excretion of lactic and carbonic acids formed during the energy metabolism, using glucose and glutamine as carbon sources. The maintenance of transmembrane ionic gradients is an important use of energy, as is cell growth. The activation of cellular receptors usually causes transient or sustained increases in acidification rate. The energetic cost of generating second messengers is probably too small to account for either change, so events more distal to the receptor-activation process must be responsible. The opening of ion channels may cause the increases in some cases. In others, changes in intracellular pH and loose coupling between ATP hydrolysis and synthesis may be involved; models for these processes are presented. Keywords: acidification, biosensor, physiometer, pH, receptor.

buffer, cell, glycolysis, metabolism,

metabolism. This is the principle of an instrument dubbed a silicon microphysiometer (Parce er al., 1989; Owicki & Parce, 1990) that we have developed, based on the light-addressable potentiometric sensor (LAPS; Hafeman et al., 1988). We have shown that when extracellular acidilication rates are measured with high precision

1. INTRODUCTION This paper examines the biological and chemical basis of biosensors that detect perturbations in the physiological state of living cells by monitoring attendant perturbations in metabolic activity. More precisely, these sensors detect the rate at which cells excrete acidic byproducts of 0965-5663/92/$05.00 0 1992 Elsevier Science Publishers

micro-

Ltd.

255

Biosensors & Bioelectronics

.I C. Owicki and .l W: Parce

Fig. 1. Activating the ml muscarinic acetylcholine receptor increases extracellular acidjicdtion rate. The rate at which Chinese hamster ovary (CHO) cells acid&d their environment was monitored in a silicon microphysiometer, Due to transfection. the cellsfunctionally expressed the human ml muscarinic receptor and the activation of this receptor with the acetylcholine analog carbachol increased the extracellular cellular acidt@ation rates in a dose-dependent fashion. Either I pum)or IO&O) carbachol was present during the period represented by the firred symbols. This system was studied by Owicki et al. (1989), and these data were obtained in collaboration with C. Bountra.

(within a few per cent) and time resolution (as line as 30 s) under culture conditions that maintain cell viability, it is possible to observe the functional effects of chemicals ranging from hormones and neutrotransmitters (Parce er al., 1989; Owicki ef al., 1990) to cytotoxic agents (Bruner etal., 1991a,b). Figure 1, for example, shows that the acidification rate of CHO cells transfected with the ml muscarinic acetylcholine receptor significantly increases in the presence of the stimulatory acetylcholine analog carbachol. We have dealt with some of the electrical- and mechanical-engineering issues elsewhere (Hafeman et al., 1988; Parce eful., 1989; Bousse et al., 1990). Here we concentrate on subjects of at least equal importance: the physical chemistry and cell biology of this type of instrument. 2. BIOSENSORS BASED ON CELLULAR METABOLIC RATE The biochemical reactions that define the living state are highly coupled together into an intricate network. Products of one reaction are reactants for another, enzyme cascades provide amplification, and feedback control loops abound. As a point of logic, it is clear that a perturbation of any biochemical reaction must ripple outward-with attenuation or amplification - throughout the entire network. This is suggested schematically in Fig. 2, where the activation of a receptor by ligand 256

binding generates a cascade of intracellular changes. As a practical point, whether such a perturbation can be detected at some remote biochemical site depends on the process monitored and on the precision and time resolution of the sensor. Intermediary metabolism, particularly the catabolic processes that yield the universal cellular energy currency adenosine triphosphate (ATP), are particularly richly coupled into the biochemical network. Therefore, metabolic (catabolic) rate is especially sensitive to the physiological state of the cell. Metabolic rate can be defined in various ways, as is suggested in Fig. 2. For example, one can monitor the rate of uptake of reactants such as glucose or 02 (Karube er al., 1982; Li er al., 1988), or the rate of production of products such as heat (James, 1987; Hammerstedt & Lovrien, 1983) or the acidic products of metabolism, lactic acid and COz (Parce etal., 1989). In somewhat related work, Rawson et al. (1989) have monitored the photosynthetic activity of cyanobacteria. We have chosen to measure extracellular acidification because of the recent availability of the LAPS, a technology that is particularly well suited for such measurements. What good is such a biosensor that monitors integrative, high-level cellular functions? Its principal strengths are two. First, it is very general, yet appropriate molecular and biological controls can be designed to assure specificity in a given experiment. If one wants to know whether the physiological state of a cell has changed in a significant way, it is probable that such a method can detect the fact. The alternative is some technique specific to the mechanism of the change; this may be difficult or, if the mechanism is unknown, impossible. The second strength, at least for the microphysiometer, is that the measurements can be practical in a laboratory setting. The system can provide an attractive combination of ease of use, time resolution, stability, and sensitivity. We now turn to the chemical and biological issues that affect the performance of an acidification-based instrument. 3. PHYSICAL-CHEMICAL

ISSUES

3.1. Introduction This section discusses how the protons produced by cells change the pH of the extracellular

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Biosensors based on the energy metabolism of living

Activated Receptpr

Ligand / Receptor

Fig. 2. Schematic representation of cellular metabolism and its relationship to physiological processes such as receptor activation. Glucose and oxygen are consumed to yield the cellular enew currency ATPand the measured waste products lactic acid and carbon dioxide. ATP is consumed by a variety of synthetic metabolic processes depicted by the array of arrows. Flux through these metabolic processes can be increased due to activation of a receptor by ligand binding. One spectfic example of an ATP-consuming reaction initiated by receptor activation is the opening of a potassium channel. This results in an e&x of potassium, lowering the cytoplasmic concentration, which in turn increases the activiv of the sodium/potassium-ATPase (depicted in the lower 1eJtquadrant of the cell).

medium, leaving questions of the mechanisms of proton production to section 4. The rate of pH change depends on the rate of proton production by cells (cell concentration and proton production per cell) and on the buffer capacity of the medium. We treat each of these in turn. 3.2. Cellular acidification rates 3.2.1. Quanttaing cells For this review we have brought together a wide variety of disparate data on extracellular acidification. To facilitate comparison, we have sought to express acidification rates consistently in terms of H+ s-l per cell. Frequently, however, absolute numbers of cells are not reported in the literature. Particularly when adherent cells are involved, it is simpler - and perfectly adequate for many purposes - instead to report a quantity directly related to the total amount of cells (e.g., the area of tissue or amount of cellular protein). To help when more detailed quantitation is necessary, we

provide the following information about cell size and composition. Mammalian cells vary tremendously in size, even within one organism. The volume of a 2 pm diameter platelet is a few femtoliters, and a large motor neuron may have an axon 1 m long by 20 pm in diameter, or 300 nl. The sizes of ‘typical mammalian cells such as fibroblasts and epithelial cells are usually in the range l-4 pl (Clegg & Gordon, 1985; Freshney, 1987, p. 233; Alberts et al., 1989, Table 3-l), on the order of 1000 times larger than an E. coli cell. When growing as a confluent monolayer, such cells often have lateral densities on the order of 10’ cells cm-*, or lo3 jf m* per cell. The protein contents of some commonly cultured cells were tabulated by Patterson (1979) and ranged between 100 and 1300 pg per cell. Significant variations occur not only between different cell types, but within a single cell type after varying lengths of culture (up to about a twofold difference). Alberts et al. (1989, Table 3-l) 257

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.I C. Owicki and J K Parce

estimate that mammalian mass, which corresponds 2 pl cell.

cells are 18% protein by to 400 pg protein for a

3.2.2 Typical rates of extracellular acidlfkation Compared to the number of studies determining the uptake and production of molecules such as glucose, oxygen and lactate, there are relatively few precise measurements of extracellular acidification rate. Balaban & Bader (1984) used a pHstat to determine that Ehrlich ascites cells and transformed hamster brain cells (HTcBH) in suspension produced 4-O X lo* and 3.6 X 10’ H+ s-l ng-’ protein with 10 mM glucose as carbon source. Acidification rates were five- to tenfold less in control medium without glucose. Given that most mammalian cells contain Ol1.0 ng protein, in the presence of glucose these cells produced z 1 X lo* H+ s-’ per cell. Parce er al. (1989) using a microphysiometer, reported acidification rates on the same order for adherent cells such as fibroblasts. 3.3 Control of pH change by buffers 3.3.1 BufSer capacig Since cells excrete protons into buffered solutions and potentiometric methods detect pH rather than [H+] directly, it is important to establish the relationship between the number of protons n excreted into a volume V and the resulting pH change. If we assume that the medium is buffered by a weak acid HA present in total concentration A, = [HA] + [A-], with dissociation constant K, the buffer capacity /3 is given by p

E _

dWJ9 = ~ W-I

~o(PK - PW ln(lO)Atot

(1

+

lo(pK-

pH))2

This can be obtained by manipulating the Henderson-Hasselbalch equation, assuming for relationship pH = simplicity the ideal -log([H+]). When multiple buffers are present, the overall buffer capacity is simply the sum of the invidual buffer capacities calculated as above. Figure 3 shows how j3 depends on pH for a single buffer. It is symmetric about pH = pK, reaching its maximum value of ln(lO)A,,,/4 z 0*58Atot when pH = pK. Although strictly speaking the relationship between n and pH is nonlinear, it is relatively linear when the pH is near the pK. Specifically, j3 varies by no more than 10% within the range 258

Fig. 3. Buffer capacity of a weak acid or base as ahnction of pH See section 3.3.1 for a discussion.

1pH - pK I< 0.28. Far away from the pK, where /3 M In(lO lo- IpK- PHI,j? is constant to within 10% for pH changes of up to 0.04. Being able to assume constant buffer capacity often simplifies data analysis. 3.3.2 Carbon dioxide equilibria Carbon dioxide is important biologically not only as a metabolite, but also because it is a weak acid that contributes to buffering and pH homeostasis (Thomas, 19893). The physiologically important equilibria are (Patton et al., 1989, p. 1120): (CO2)aq

+

H20

+

H2C03 --H+

+ HCOT It is usually not necessary to consider the further dissociation of bicarbonate to carbonate. Dissolved CO2 can also partition into the gas phase (if it exists), which is how terrestrial animals dispose of CO,; its solubility, considering only (C02)aq, is about 0.03 mM per torr partial pressure of CO2 at 310 K Under physiological conditions the equilibrium strongly favors dissociation (pK = 6.1 for [H+] [HC0,1/[C02] at 310 K and physiological ionic strength). In mammals, and under most cellculture conditions, the aqueous CO2 is in equilibrium with a local atmosphere containing about 5% C02. This implies that at physiological pH of 7.4, [HCOJ = 24 mM and [CO,] = l-2 mM. The buffer capacity due to these species is 2.6 mM PH-‘, assuming a closed system. If the CO2 can escape into the atmosphere, the buffer capacity is instead given by log(e) [HCOJ, or 10 mM. In either case, this is an important contribution to the buffer capacity of the extracellular fluid. It dominates the buffer capacity of most standard

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Biosensors based on the energy metabolism of living cells

mammalian tissue-culture media. Hence, if there are no adverse physiological consequences it is often advantageous to increase the sensitivity of acidification measurements by not adding CO2 and NaHC03 to the media. This also eliminates issues of pH drift due to the gas permeability of containers and tubing. Regardless of whether CO2 is intentionally added to the medium, it will be produced metabolically by most cells. In a closed system near physiological pH each CO2 will produce nearly 1 H+ at equilibrium. However, the hydration step can be a kinetic bottleneck: the half-life for a dissolved COZ is about 11 s at 3 10 K and pH 7.4. This is calculated from a combined pseudo-first-order rate constant of 0.062 s-t for hydration based on the data of Pinsent etal. (1956) for reaction of CO2 with HZ0 and OH-. Hence, not all the CO* produced may be counted during rapid measurements of acidification. The slow hydration of CO* presents physiological problems for CO? transport in metazoa; carbonic anhydrase catalyzes the hydration very efficiently and is present in the cytoplasm of many types of cells (Tashian, 1989). This enzyme can be included in the extracellular medium in an instrument. A simple model shows the effects of CO2 hydration on measured pH changes. Assume that CO* is produced at a constant rate, that it is hydrated by a first-order process with rate constant khyd and then instantly dissociates to H+ and HCOF, and that the buffer capacity of the medium is constant. This is a system of two linear differential equations that can be solved to give the following expression for the pH change in time r = khyd X t for the uncatalyzed case compared to that for the fully catalyzed case (k hyd = a):

mechanistic information: it should be possible to differentiate between lactate and COZ production by comparing catalyzed and uncatalyzed aciditication rates. This possibility strictly depends on being able to measure acidification rates rapidly.

ApH,,,,JApH,,,

= 1 - [l - exp(-r)]/r

This same expression also gives the ratio of the mean acidification rates during the period. The ratio is 0 when t = 0,0*05 when r = 0*1,0.37 when r = 1.0,090 when t = 10, and 1 when r = 00. In preliminary experiments using the microphysiometer, enhancements of acidification rates with carbonic anhydrase have indeed been observed (D. Miller, unpublished results). It is apparent that catalysis can significantly increase acidification rate due to CO2 production when the measurement time is !Z l/khyd or less. Beyond increasing sensitivity, this can provide

3.3.3. Buffering in cell-culture media The formulas for many common cell-culture media are conveniently tablulated by Ham & McKeehan (1979). In the absence of deliberately added COJHCOT or a buffer such as HEPES (pK = 7.3 1 at 310 K), the dominant buffer at physiological pH in most media is phosphate. This is typically present at z 1 mM, with RPM11640 medium having a notably high level (5.6 mM). At 3 10 K the second pK for phosphate is 6.76 (Dawson et al., 1986). Thus, at pH 74O,p/A,, = 0.35 for phosphate. This neglects the possible presence of M2+, which removes an MgHP04 complex (Kd = 2 mM; Phillips ef al., 1969) from the proton equilibrium and would therefore reduce the buffer capacity. Amino acids are typically present at an aggregate concentration of 5-10 mM and may contribute zz 0.2-0.3 mM to the buffer capacity, mostly due to their a-amino groups. The amino acids with highest intrinsic buffer capacity at pH 7.4 are histidine and cysteine, but glutamine is usually the largest contributor to the buffer capacity of the medium because it is present in the highest amount. The buffer capacity of a typical medium, then, is a little less than 1 mM. Serum is often present at levels from 1% to 15% (v/v). We have measured the buffer capacity of fetal bovine serum at 310 K, pH 7.4, and ambient CO2 levels to be 7 mM pH-‘. Hence, the buffer capacity of the added serum is roughly comparable to that of the medium. 3.4. Minimum cell concentration for a biosensor The concentration of cells should be high enough that biological acidification rates are at least ten times the statistical uncertainty in acidificationrate measurements. We estimate the latter at about 30ppH s-l for measurements on the minute time scale in the present version of the silicon microphysiometer. If a cell produces lo* H+ s-t, or 2 X lo-l6 mol H+ s-t, then with 1 mM pH-’ buffer capacity the concentration of cells should be at least 1 X lo6 cells ml-‘. This is generally easy to achieve; 1 ml of packed cells usually contains lo*-lo9 cells.

.I C. Owicki and J W Parce

Note that since the measurement is potentiometric it is the concentration and not the number of cells that is fundamental. Miniaturization of the instrument is advantageous from several viewpoints, and the minimum size is limited by mechanical- and electricalengineering constraints.

4. CELL-BIOLOGY

ISSUES

4.1 Introduction Our previous studies (Parce etal., 1989; Owicki er al., 1990) were well controlled but largely phenomenological. For the approach to be most useful, it will be necessary to understand the underlying cell biology better. In this paper we cull information from the existing biochemical and cell-biology literature with two purposes. First, we try to define the biological phenomena most relevant to extracellular acidification. Second, where possible we attempt to make quantitative estimates in terms of effects on extracellular acidification rate. Both should illuminate existing experiments and suggest new experiments to optimize and better understand this class of biosensor. We are not attempting to make a critical review of such a broad field of biology, but instead to recast the literature in a new context. Our scope is further limited primarily to mammalian cells, although most of the principles apply to the cells of other animals and also substantially to those of plants, fungi, protists, and bacteria. We particularly exclude a substantial literature in which cells, especially bacteria, are used only as catalysts to transform an analyte into a form that can be detected conveniently. Even within this limitation, and even given the underlying biochemical unity of life, there is a great deal of biological diversity: among organisms, among tissues, and even among the same kinds of cells under different growth conditions. The reader should keep this in mind while reading generalizations about ‘typical’ cells. In section 4.2 below we describe the most important metabolic processes that produce acid, as well as the principal mechanisms by which the acid leaves the cells. Changing the focus from energy supply to energy demand, in section 4.3 we discuss aspects of cell physiology that determine

Biosensors & Bioelectronics

cellular metabolic acidification.

rate

and

extracellular

4.2 The production and excretion of metabolic acid 4.2.1 Introduction For detailed analysis of the pathways discussed here, see standard biochemistry texts such as Lehninger (1975) or Stryer (1988), or reviews of the metabolism of cultured cells such as Paul (1965) and Morgan & Faik (1981). The primary carbon sources for metabolic energy are sugars, amino acids, and fatty acids. In culture, glucose and glutamine are usually present at high (mM) levels in the culture medium; fatty acids are seldom added except to satisfy specific biosynthetic requirements. Table 1 summarizes the ATP-yielding reactions that we discuss, including the ATP per H+ produced. We have also assumed that the reactions are as efficient as possible. If oxidative phosphorylation is partially uncoupled or if futile cycles are active (e.g., involving hexokinase and glucose-6-phosphatase (Lehninger, 1975, p. 631)), then more protons will be produced per ATP. ATP does not appear explicitly in the chemical equations in Table 1, because we have assumed that all the ATP produced is then hydrolyzed so that there is no net change in the ATP concentration in the cell. As has been pointed out by Gevers (1977) and by Hochachka & Mommsen (1983) ATP synthesis per se consumes protons due to the differences in protonation states of the reacting species: (MgADP] + (Pi) +yH+-+

(MgATP) + Hz0

Here the notation ( ) indicates all ionization states and degrees of complexation with M$+. At pH 6.8,~ = 0425; at pH 7.4,~ = @8; and at pH 8.00, y = 0.945; all are calculated for 4.4 mM free M$‘, a value characteristic of muscle. For [M$+] = 1 mM, perhaps more typical of most eukaryotic cells, y would be reduced by 0.1-0.2 (Phillips et al., 1969). ATP hydrolysis simply reverses the above reaction. The assumption of tight coupling between the rates of ATP synthesis and hydrolysis in Table 1 will be examined later in the paper. 4.2.2. Glucose metabolism Glucose can be converted to lactic acid via glycolysis, or to CO2 via respiration (glycolysis

Biosensors based on the energy metabolism of living cells

Biosensors & Bioelectronics

TABLE 1 Summary of principal energy-yielding

metabolic pathways

Carbon Source

Pathway

Reaction”

ATP yield

Glucose Glucose Glucose

glucose + 2 lactate- + 2H+ glucose + 602 + 6HCO: + 6H+ 3 glucose + 11/202 + 5 pyruvate+ 3HC0, + 5H20 + 8H+

2 36b 27d

1@00 0,167 0,296

Glutamine

Glycolysis Respiration HMP shunt + glycolysis + oxidative phosphorylationc Respiration’

27

0.111

Pyruvate

Respiration

glutamine + 9/20, + 3H,O + SHCO, +2NH;+3H+ pyruvate- + 5/202 + HZ0 + 3HC0, + 2H+ Cz,, HZn_ ,O, + (6n - 2)02 --+ 2n HCO, + (2n + l)H+

15

0.133

17 n-6

Q129(n = 9)

Fatty acid /3-Oxidation, respiration

H+ per ATP

“Assumes that all the ATP produced is hydrolyzed and all the CO1 produced hydrates and dissociates into H+ and HCO,. At pH 7.4,95% of the CO2 does so (pK = 6.1). bAssumes that each cytosolic NADH produced in the glycolytic part of the pathway yields 2 ATP. If a more efficient shuttle system is used to pass the NADH to the mitochondrion the yield rises to 3 ATP and the overall yield of ATP for the reaction becomes 38. ‘The HMP shunt alone produces no ATP; see text. dAssumes 2ATP per NADH or NADPH: see footnote b above. =See text for pathway.

followed by the citric acid cycle and oxidative phosphorylation). As Table 1 shows, not only is respiration more energetically efficient than glycolysis, but it produces much less acid for each ATP generated. Glycolysis produces far more H+ per ATP synthesized and hydrolyzed than any other major pathway. Although respiration is the major source of ATP from glucose under most conditions in viva, glycolysis usually predominates in vitro. Exceptions exist; see, for example, Kemp eral. (1990). The high glycolytic activity of cells in culture is due partly to the fact that many cultured cells are tumor lines, which have been known since the work of Warburg (1926) to be highly glycolytic. Another contribution is more intrinsic to the culture conditions. When cells are removed from an organism and placed in culture, the metabolic balance often tips from respiration toward glycolysis, sometimes on the time scale of hours (Gebhardt etal., 1978). The cause of this phenomenon is not well established, but might be related to hypoxia in vitro (Mandel, 1986). Production of CO2 from glucose does not necessarily demonstrate respiration, since the CO2 is often produced by the hexose monophosphate shunt in vitro (Morgan & Faik, 198 1). The shunt normally is not a major source of ATP, but instead provides NADPH for

biosynthesis. However, if pathways are present to couple NADPH to oxidative phosphorylation (perhaps by transhydrogenase; see Lehninger (1975, p. 499)), the shunt can be used to produce energy. Table 1 shows one scheme that is consistent with the observation (Morgan St Faik, 1981) that the CO* generated in cell culture is often predominantly formed from Cl of glucose if glucose is the only carbon source. The reaction shown is essentially that described by Stryer (1988, p. 433). 4.2.3. Glutamine metabolism It has been estimated that mammalian cells in culture obtain 30-100% of their ATP from the oxidative catabolism of glutamine, depending on cell type and the hexose content of the medium (Zielke ef al., 1984). The metabolic pathways for glutamine degradation that are listed in Table 1 are consistent with the observed products: glutamate, aspartate, CO*, and NH3. The production of NH3 suggests that glutamine metabolism might result in a consumption rather than a production of H+. Not so. Neither of the successive deaminations by glutaminase and glutamate dehydrogenase to produce a-keto-glutarate produces or consumes a proton if ATP hydrolysis is tightly coupled to synthesis. The oxidation of glutamine is 261

.I C. Owickiand J. WVParce

completed when a-keto-glutarate enters the TCA cycle, exits as oxaloacetate and re-enters as acetylCoA via conversion to phosphoenolpyruvate and then pyruvate. The overall oxidative degradation of glutamine causes a small net acidification, 1 H+ per 9 ATP formed and hydrolyzed. Aspartate can be produced from some of the intermediates in glutamine degradation in a transamination reaction that does not in itself affect acidity (glutamate + oxaloacetate + aspartate + a-ketoglutarate). 4.2.4. Other carbon sources Table 1 lists energy-yielding reactions of two other important metabolites: pyruvate and fatty acids. Pyruvate can enter directly into respiration, as can fatty acids after degradation to acetyl-CoA by @-oxidation. Cells need not depend entirely on the import of carbon sources for the production of ATP. Glucose is stored in the cytoplasm in the polymer and fatty acids are stored as glycogen, triglycerides. The degradation of glucose formed from glycogen is similar to that of exogenous glucose, except that one more ATP per glucose is produced in glycogenolysis (no ATP is necessary to generate the glycolytic intermediate glucosedphosphate). The hydrolysis of fat to glycerol and three fatty acids neither consumes nor produces ATP, but it does produce one H+ per fatty acid due to the dissociation of the carboxyl group. The amounts of these storage molecules depend on cell type. For example, liver and muscle cells have large stores of glycogen, and adipocytes contain large amounts of triglycerides. Instead of storing only the carbon source, some cells can also store high-energy phosphate bonds. The most common form is phosphocreatine, which can deliver high power in skeletal muscle cells and may be present there at concentrations five times that of ATP (Lehninger, 1975, p. 767). It also occurs in nerve and smooth-muscle cells, but not appreciably in many other mammalian cells and not in bacteria (Lehninger, 1975, p. 409). The equation for the donation of the phosphate group of phosphocreatine to ADP to form ATP, plus the subsequent hydrolysis of ATP is (Hochachka & Mommsen, 1983): (creatinep)

+ Hz0 + 0*15H+ + creatine + Ipi)

before, () indicate a sum over the different ionization states. At pH 7.4, for which the

As

262

Biosensors& Bioelectronics

reaction is written, the hydrolysis consumes a modest amount of H+. At pH 6.8, 0*42H+ are consumed, and at pH 8.0, only O@H+. Thus, a burst of energy use in the form of the hydrolysis of phosphocreatine would tend to increase rather than decrease pH. This is opposite to the effect for ATP hydrolysis. 4.2.5. Acidproductionji-om specialized metabolism Some cells produce substantial amounts of extracellular acid as part of specialized secretory activities. This may exceed the rate of extracellular acidification due to conventional metabolism. We briefly discuss three examples here: the respiratory burst in neutrophils, lipolysis in adipocytes, and proton transport in epithelia. When phagocytosis is stimulated in neutrophils there is a burst of 02 consumption due to the activation of the hexose monophosphate shunt. The purpose of this is thought to be the formation of antimicrobial 0, via NADPH oxidase. Borregaard et al. (1984) have used a pH-stat to study the extracellular acidification of = lo* neutrophils in 10 ml before and during the respiratory burst. They found that resting neutrophils produced 1.3 X lo7 H+ s-’ per cell, a rate that increased more than fourfold to 5.5 X lo7 s-t per cell during a respiratory burst that lasted several minutes. The resting cells produced no detectable COz, and all the extra protons produced by the stimulated cells could be accounted for by the CO2 produced during the respiratory burst, presumably by the hexose monophosphate shunt. Adipocytes are fat-storage cells whose large size is attributed to the presence of a droplet of triglycerides that may occupy more than 90% of the cell volume. The stored fat is mobilized for use by other tissues by hydrolysis to glycerol and fatty acids, which are transported across the plasma membrane. Since fatty acids have pKs substantially below physiological pH, lipolysis produces one H+ for each fatty acid. Nilsson & Belfrage (1979) analyzed extracellular acidification rates of rat adipocytes (100~1 packed cells in 2 ml, 1.6 X lo6 cells) using a pH-stat. Acidification rates in the absence of lipolysis were difficult to detect but rose to 1.4 X lo9 H+ s-’ per cell at maximum stimulation with norepinephrine, a rate sustained for 30 min. Epithelia are interfaces between different biological compartments, such as the blood and the lumen of the gut. Epithelial cells are usually

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Biosensors based on the energy metabolism of living cells

polarized, i.e., the plasma membranes that face the two compartments are st~c~rally and functionally different. An important function of such cells is the transport of ions and molecules from one compartment to the other. A familiar example is the absorption of nutrients from the gut, but proton transport can also be important. For example, Weinman & Reuss (1982) used extracellular pH measurements to study proton transport through the apical membrane of a gallbladder epithelium. The parietal cells of gastric glands secrete HCl at PI-I < 1, and the proximal and distal tubules of the kidney are major sites of pH regulation of the blood. The common mechanism of proton transport involves proton extrusion into the lumen of the organ via the HSf K+ ATPase in the apical membrane with maintenance of intracellular pH via Cl-/HCOF exchange through the basolateral membrane into the blood (Patton mal., 1989, p. 1091 ff.). Helander (1977) has estimated that each parietal cell in the rat transports 7 X IO* H+ s-’ into the stomach, which is considerably higher than the basal acidification rate of most cells. Essentially this same mechanism acts in a nonepithelial cell, the osteoclast. Osteoclasts tunnel through bone in part by secreting protons that dissolve the mineral in small contact regions (Baron, 1989). For any of these proton transport processes to be studied as extracellular acidification in vitru, it is desirable to preserve some semblance of the polarized state that is associated with epithelial architecture (or osteodast-bone contact) in vivo.

protons that are coupled to other sources of free energy. For reviews and references, see Roos & Boron (1981) and Andersen (1989). An important motif is the exchange of a cytoplasmic proton for an extracellular cation. The plasma membranes of mammalian cells commonly contain a protein that catalyzes the Na+/H+ exchange. Since this is electrically neutral and the concentration gradient of Na+ exceeds that of H+, the result is an excretion of H+; the influx of Naf is reversed by the Na+/K+ATPase at the expense of ATP. In acid-excreting epithelia, H+/K+ exchange (against both ionic concentration gradients) is directly coupled to ATP hydrolysis. Alternatively, there are H+ATPases that transport protons without cotransport of other ions. These examples involve the direct transport of a proton. Essentially the same effect can be achieved indirectly, by transposing a weak acid, which produces a proton upon dissociation after transport; weak bases have the opposite effect. Both uncatalyzed and catalyzed instances of this strategy are physiologically important. The uncatalyzed case is based on the high permeability of membranes to small uncharged molecules such as CO2 and NH3, but not their charged counterparts HCOT and NH:. One consequence is that the metabolic production of CO2 need not lead to intracellular acidification. The transmembrane transport of the physiologically important weak base HCOS is facilitated by several mechanisms involving cotransport of other ions. The best known is the band 3 Cl-/HCO, exchange protein from erythrocytes, which permits the efticient transport of respiratory CO2 from tissues to the lungs. Cl-/HCO; exchange as well as mechanisms such as Na+/HCO: co-transport operate in other tissues as well. Given the importance of glycolysis for cultured cells, the transport of lactate and lactic acid across the plasma membrane is particularly significant for pH and energetic homeostasis in culture. However, it has been studied most thoroughly in three systems for which it is particularly irn~~nt physiolo~~ally: muscle, liver, and erythrocytes (reviewed by Juel (1988) and Deuticke (1989)). Multiple mechanisms exist, including facilitated transport by monocarboxylate carriers and anion exchange proteins, as well as unfacilitated diffusion of the protonated acid across the membrane.

Me~bolism generally produces protons at a rate faster than they could diffuse passively across the plasma membrane, which is not very permeable to ions (including protons) except via specific ion channel and carrier proteins. Although proton channels do exist (Thomas, 1989a), they cannot be the primary route for the disposal of excess protons, since the electrochemical gradient for protons in most cells would lead to an inward flux; cytopiasmic and extracellular pH are usually similar in mammalian cells, but the transmembrane potential is negative inward. Thus, there must be specific mechanisms for excreting

263

J C. Chvicki and J. W Parce

4.3. Factors that influence the rate of metabolism 4.3.1. The regulation of metabolic rate The production and consumption of metabolic energy are linked by a variety of intricate mechanisms, the best studied of which involve the balance between ATP and its precursors ADP and AMP (Lehninger, 1975, p. 537). A preponderance of ATP generally indicates that the cell is in an energy-rich state, while the reverse indicates the need to increase metabolic rate. Mass-action effects of ATP and ADP are involved in controlling the rate of ATP synthesis in the mitochondrion, though other factors probably also are important (Balaban, 1990). ATP and ADP are also allosteric modifiers of key enzymes in glycolysis and the TCA cycle, ATP inhibiting and ADP enhancing activity. An example is phosphofructokinase-1 (PFK-I), a critical control point in glycolysis that catalyzes the phosphorylation of fructose-6-phosphate to fructose-1,6-bisphosphate. This enzyme is also subject to an entirely different kind of control that is related to receptor activation rather than cellular energy charge. Its activity is stimulated allosterically by fructose-2,6-bisphosphate (fru2,6-P2). In some tissues the activity of enzymes responsible for the formation and degradation of this modulator are controlled by CAMPdependent kinases, which renders fru-2,6-P2 effectively a third messenger for ligand-receptor systems that act by altering CAMP concentrations (Hue & Rider, 1987). Yet another source of metabolic regulation is changes in intracellular pH (pHii Roos & Boron, 1981; Busa &Nuccitelli, 1984, Busa, 1986). Fertilization, the activation of growth-factor receptors, and a variety of other activation phenomena lead to cytoplasmic alkalinizations on the order of tenths of a pH unit. In some tissues the rate of glycolysis increases dramatically at slightly elevated pH, due in part to increases in PFK-1 activity (e.g., Fidelman et al., 1982). Glycolytic control is not confined solely to PFK-1. Other enzymes participate, as does the glucose transporter. For example, insulin can increase ability of a cell to transport glucose into the cytoplasm, perhaps by mobilizing an intracellular reservoir of glucose transporters into the plasma membrane (Saltiel, 1990). Thus, modulation of PFK-1 activity by fru-2,6Pz and by intracellular pH provides direct links between receptor activation and increased extra264

Biosensors & Bioelectronics

cellular acidification. This is mediated by an increased rate of glycolysis and, under circumstances discussed below, by the protons excreted from the cells to increase pHi. 4.3.2. The relative importance of various physiological finctions 4.3.2.1. Cell growth: biomass. The rate at which cells acidify their environments is the product of two factors: the number of cells present and the rate of acid excretion by a typical single cell. In this section we primarily discuss cell number, turning to the rate per cell in subsequent sections. The acidification of culture medium by cells is a common experience of those who culture cells, and it has long been used semi-quantitatively to detect the viability and growth of cultured cells. For example, cells in a microplate are exposed to a virus or cytotoxic compound (Paul, 1975). After a period allowing for cell growth - typically 2-14 days - the color of the medium is noted. Culture medium usually contains the pH indicator phenol red (pK = 7*9),which changes from red to yellow as the pH drops. A yellow color at the end of the incubation indicates a high cumulative metabolic activity, while a pink color indicates cytotoxicity and/or growth inhibition. Rather than monitor the accumulation of acid over such long periods, we have measured cellular acidification rates in the silicon microphysiometer, a flow system in which the culture medium is continually refreshed. When other biochemical factors are held constant, the acidification rate is proportional to cell number and can be shown to increase exponentially with time when microbial cells are in log growth phase (J. Libby and D. Miller, personal communication). Using the same system to monitor decreases in aciditication rates, it is possible to measure cytotoxicity more quantitatively and with greater time resolution than in the microplate system. For example, inhibition of extracellular acidification on the time scale of minutes was found by Bruner et al. (1991b) to correlate well with the ocular irritancy of consumer products. Such measurements include contributions both from a decreased number of viable cells and from lowered metabolic rates throughout the population. On a longer time scale, cytopathic effects of a slow-acting virus (HIV) and efficacy of the antiviral agent AZT were determined over six days of continuous culture in the microphysio-

Biosensors & Bioelectronics

Biosensors based on the energy metabolism of living cells

meter (Wada etal., 1991b). Since the doubling time of a mammalian cell line is usually about a day, experiments on this time scale strongly reflect cell growth. On an intermediate time scale, a few hours to a day, we have monitored chemotherapeutic efficacy on tumor cells (Parce etal., 1989), cytopathic effects of a rapidly acting virus (vesicular stomatitis virus) and protection by an antiviral agent (Parce etal., 1989) and the detection of the cytokine interleukin-2 (Fok & Wada, 1991).

energy costs of the process. For example, fairly actively secreting hybridomas may produce about 1 X lo3 IgG s-’ per cell (Schneider & Lavoix, 1990). A lower bound for the biosynthetic costs is three ATP molecules hydrolyzed per peptide bond formed (Paul, 1965). A 150 kDa IgG molecule has about 1300 peptides, so this corresponds to 4 X lo6 ATP s-r per cell. Miller et al. (1989) have determined that a hybridoma (albeit a different line under nonidentical culture conditions) uses 2 X lo8 ATP s-’ per cell overall, so it appears that at least a few per cent of the metabolic energy of such hybridomas is expended on secreted protein. This should be detectable by current methods. Whether the proportion is higher depends on factors such as metabolic efficiency and the costs of any synthesis of amino acids that are not obtained from the medium, as well as membrane transport. We are aware of no direct experimental determinations of secretory energy costs.

4.3.2.2. Cell growth: metabolic costs. What fraction of a cell’s metabolic energy is dedicated to growth, as opposed to mere maintenance activities? This question has particularly been studied by biochemical engineers for fermentation systems. For example, murine fibroblastic LS cells grown in suspension were found by Kilburn et al. (1969) to use 65% of their ATP for maintenance and 35% for growth. Similar results (60% and 40%) were found for hybridomas by Miller et al. (1989). Skog et al. (1982) determined that about one-third of the ATP produced by Ehrlich ascites cells was consumed by protein synthesis, and about 1% by nucleic acid synthesis. Kilbum et al. (1969) noted that the amount of ATP consumed in producing a given dry weight of LS cells was similar to that for a bacterium and a mold, but the fraction of that ATP that was expended on maintenance was much higher in the LS cells.Aerobacteraerogenes was calculated to expend only 7% of its ATP on maintenance. These differences are largely due to the slow mammalian cell cycle, which necessarily enhances the comparative importance of maintenance activities. Note that the results quoted above do not necessarily predict the differences in energy requirements between rapidly dividing and nonmitotic cells. For example, cell growth can be arrested by removal of growth factors (serum starvation). This leads to profound physiological changes in the cells, beyond merely affecting growth-related activities. When growth arrest is removed, the metabolic rates can increase several-fold, as has been shown for 3T3 Bbroblasts by Diamond et al. (1978). 4.3.2.3. Protein secretion. Beyond growth and maintenance, some cells have another biosynthetic load: the synthesis and secretion of proteins or other molecules. It is instructive to estimate the

4.3.2.4. Maintenance of ion gradients. The maintenance of ionic gradients across the plasma membrane and the resulting transmembrane electrical potential is essential. The cytoplasm of a typical mammalian cell contains 5-15 mM Na+ and 140 mM K+, while the extracellular concentrations of these ions are essentially reversed (145 mM and 5 mM); the electrostatic potential of the cytoplasm is about -20 to -200 mV with respect to the extracellular medium (Alberts et al ., 1989, p. 316). The plasma membrane is slightly permeable to both cations, so energy must be expended to maintain the nonequilibrium distribution. The plasma membrane of essentially all animal cells contains an integral membrane protein that functions as an ATP-driven ion pump, catalyzing the following reaction (section 4.2.1 and Alberts et al., 1989, p. 305): 3CNa+)wo + 2(K+),fim + lMgATP1+

H2G

+ 3(Na+)eti,a + 2(K+)Cy,o + (MgADP) + {Pi) + yH+ Note that beyond maintenance of the Na+/K+ gradient there is a net transport of one cation out of the cell, which contributes to the membrane potential and is an important osmotic mechanism for regulating cell volume. This system is typically a major sink of metabolic energy. The brain devotes >50% of its 265

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JV C. Owickiand .I W Parce

metabolic energy to the Na+/K+-ATPase (Lehninger, 1975, p. 838). For human erythrocytes the fraction is 20% (Wittam &Alger, 1965), and for transformed hamster brain cells HTcBH 30% (Balaban & Bader, 1984). For Ehrlich ascites cells estimates of 30% (Balaban & Bader, 1984) and 50% (Skog et al., 1982) have been obtained. In a particularly interesting study, Harris et al. (1981) measured the rate of oxygen consumption of freshly excised proximal tubules from rabbit kidney as well as mitochondria isolated from this preparation. Under normal conditions, the cells respired at 56% of the maximum rate that their mitochondria were able to sustain. When the Na+/K+-ATPase was inhibited with ouabain, the rate fell to 30% of the maximum, and when the pump was fully activated by permeabilizing the membrane to Na+ and K+ with nystatin, the rate rose to 100% of the maximum. These data imply that under normal conditions nearly half of the cells’ ATP was hydrolyzed by the Na+/K+ATPase. Under extreme stress, the ATP hydrolyzed by the pump was more than double that due to all other processes. 4.3.2.5. Motion. The generation of motion, as in muscle cells or cellular motility, is directly coupled to ATP hydrolysis. For example, oxygen consumption by human skeletal muscle is more than 20-fold higher during heavy work than rest (Lehninger, 1975, p. 832). On top of this, heavy TABLE 2 Receptors whose activation

work causes a large increase in the rate of glycolysis. The cytoskeleton of non-muscle cells may also be a significant consumer of ATP. For example, Tillmann & Bereiter-Hahn (1986) found that disrupting actin filaments with cytochaslasin B in endothelial cells reduced energy consumption by 25%. The fraction of cellular ATP expended on cell motility varies widely with cell type. Hammerstedt et al. (1988) have estimated that 60% of the ATP in bull sperm is used for motility. In contrast, Jauker et al. (1986) estimate that only about 1% of the ATP use by the motile protist Tetrahymena is used for ciliary motion.

4.3.2.6. Receptor activation. 4.3.2.6.1. Experimental observations. As noted earlier, we have observed that pharmacologically relevant changes in extracellular acidification with the silicon rates can be detected microphysiometer when a wide variety of cellular receptors are activated. These are summarized in Table 2. The kinetic profiles of the responses differ. Although it can be simpler or more complicated, the response is commonly a transient, usually an overshoot, that settles down to a new steady state characteristic of the new physiological state in the presence of the ligand. Similar results have been obtained in a few other cases using glass electrodes and larger quantities

has been detected in the silicon microphysiometer

Superfamily if known

ml-Muscarinic acetylcholine Muscarinic, subtype unknown &-Adrenergic Prostaglandin (E) Epidexmal growth factor

G protein, inositol phosphate G protein G protein, increase CAMP G protein, increase CAMP Growth factor, tyrosine kinase

Insulin, insulin-like growth factor Glutamate (kainate)

Growth factor, tyrosine kinase

.

Excitatory amino acid, ion channel

.

y-Interferon

Hematopoietin

superfamily

.

Interleukin-2

Hematopoietin

superfamily

.

Interleukin-4

Hematopoietin

superfamily

T-cell

T-cell receptor superfamily

266

and mechanism,

Native

Receptor

Transfected .

. . . .

.

Reference Owicki et al. (1990) Miller et al. (1991) Owicki et al. (1990) Parce et al. (1990) Parce et al. (1989X Owicki et al. (1990) Rice d al. (1991) Raley-Susman et al. (1990) Indelicate et al. (1991) Fok & Wada (1991) Wada et a[. (1991b) Indelicate et al. (191) Wada etal. (1991~)

Biosensors & Bioelectronics

of cells, for example the activation of platelet aggregation by thrombin (Akkerman & Holmsen, 1981). These responses are significant in at least two ways. They provide a new way to observe some of the physiological consequences of receptor activation, and they provide a general means relatively independent of the ligand, receptor, transduction mechanism, and cell type - for detecting receptor activation or its inhibition. We devote the next section of the paper to a more detailed discussion of the sources of changes of extracellular acidification rate when the cellular physiological state changes, for example by receptor activation. 4.3.2.6.2. Energy consumption due to the transduction mechanism. As discussed earlier, a typical rate of acid production for cultured cells is - lo8 H+ s-’ per cell. The ATP production rate must be on the same order of magnitude for these primarily glycolytic cells. Because triggering a receptor usually increases the acidification rate from 10% to 100% of the resting rate, the increase in the rate of synthesis of ATP must be on the order of lo7 to 10’ ATP s-r per cell. Can these changes in extracellular acidification rate either transient or sustained - simply reflect the biosynthetic costs of production of second messengers such as CAMP or phosphatidyl inositols and diacyl glycerol, or the activity of protein kinases? Although little literature on this subject is available, it is possible to make some estimates of the amount of ATP consumed in generating second messengers. Consider CAMP production induced by adrenergic agonists in tibroblasts transfected with/3-adrenergic receptors. The concentration of intracellular CAMP increases at a rate of 50 pm01 (Chung et al., 1988), min-’ mg-’ protein consuming Z 10’ ATP s-’ per cell. The actual flux of ATP through the CAMP pathway may be considerably higher than the rate at which the concentration of CAMP rises within the cell, as shown by 180 labeling studies using parotid gland tissue (Deeget al., 1988). This high flux would still account for only z lo6 ATP s-l per cell. Based on these calculations, we would expect to see only a M 1% increase in the cellular acidification rate due to the increased energy demand required to synthesize CAMP in response to receptor activation. For receptor-activated kinase activity, little

Biosensors based on the energy metabolism of living cells

quantitative information is available. Assuming that the average cell protein has a molecular weight of 50 000, consumption of lo8 ATP s-l per cell by the kinase activity would phosphorylate 20% of the entire cellular protein every second. Clearly, the ATP spend on protein phosphorylation cannot account for the observed receptormediated metabolic bursts. A similar argument can be made for phosphatidyl inositol hydrolysis and resynthesis. Some caution must be used in interpreting this type of calculation, however, because it is the fluxes through these pathways rather than concentration changes that determine their contributions to the rate of ATP consumption; as indicated above, few measurements of such fluxes have been made. In general, the flux through second messenger pathways is probably not more than an order of magnitude greater than the rate of change in steady-state concentrations of the messengers seen within the cells. Therefore energy consumption by second messenger pathways probably represents a small percentage of basal acidification rates and would not explain a substantial part of the much larger changes in rates that are usually seen during receptor activation. The opening of ion channels as a result of receptor triggering, on the other hand, may account for a considerably larger fraction of the total cellular ATP flux. Whole-cell patch-clamp experiments on A9L cells transfected with the ml muscarinic receptor have demonstrated potassium currents of M lo9 K+ s-’ per cell in response to agonist (Jones etal., 1988). This would require about half that rate of ATP consumption in order for the Na+/K+-ATPase to maintain ionic homeostasis. These data can be misleading in that the K+ flux out of non-voltage-clamped cells will hyperpolarize the cells and limit further K+ flow through the channels. Rates on the order of lo8 K+ s-’ per cell can be calculated from data based on the release of potassium to the extracellular medium from parotid gland tissue in response to receptor agonist administration (Butcher et al., 1976). Even without voltage clamping, the ATP consumption rate for this example is still comparable to basal ATP production rates. Although the energy required to produce second messengers does not appear to play a major role in the metabolic burst caused by receptor activation, in some cases ion pumping required to maintain homeostasis in the presence 267

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.I C. Gwicki and J. W: Parce

of open ion channels appears to account for consumption of ATP commensurate with the metabolic burst. 4.3.2.6.3. Models for transient behavior during receptor activation. One source of transients such as those seen in Fig. 1 and in the studies cited in Table 2 may be imperfect coupling between ATP synthesis and hydrolysis. The tight coupling assumed in Table 1 must be true on the average over long periods. However, when the rate of ATP hydrolysis changes very rapidly there is a time lag before the rate of synthesis adjusts to maintain a steady-state ATP concentration. In this section we discuss the time scale of this transient change in ATP concentration and its consequences for extracellular aciditication. Recall that the production of ATP, without considering its hydrolysis, is less acidifying than is implied in Table 1. For example, the synthesis of one molecule of ATP via glycolysis yields only (1 -Y) = O-3 protons instead of 1. Subsequent hydrolysis of the ATP generates the remaining y z 0.7 protons. Synthesis of ATP from glutamine by respiration actually consumes protons, yielding (2 - 15y)/15 z -0.57 protons per ATP. Again, ATP hydrolysis yields the value of 0.13 protons per ATP that is shown in Table 1. The effects of loose coupling depend on the metabolic pathways active in the cells. Consider an instantaneous 30% increase in the rate of ATP hydrolysis. For a cell deriving its ATP purely from glycolysis, the instantaneous percentage increase in extracellular acidification rate is (0.3 X y/l) X 100% N 21%. For a cell deriving its ATP from glutamine respiration, the increase is much larger: (O-3 X Y/0.111) X 100% = 189%. Realistically, rates of change ofATP hydrolysis are not instantaneous, and ATP hydrolysis is coupled to ATP synthesis by feedback mechanisms. The results of one simple model are shown in Fig. 4. Note that under some conditions, e.g. oxidative metabolism of glutamine or glucose, the increased demand for ATP causes a transient overshoot in the extracellular acidification rate. What about the time scale of the transients? A reasonable estimate is the turnover time for cellular ATP; this is in fact accurate for the simple model in Fig. 4. In two studies on mitotically synchronized cell populations the ATP turnover time was found to vary over the cell cycle within the rage 15-19 s for L tibroblastic cells (Ishiguro 268

1.4,

4

Fig. 4. Modelfor the effects of loose coupling between ATP synthesis and hydrolysis on the extracellular acidification rate. In the top panel, the rate of ATP hydrolysis (long dashed line) is smoothly increased as indicated, perhaps in response to the activation of a receptor. It is assumed that ATP levels are regulated by a simplefeedback mechanism: noncompetitive inhibition of a key synthetic enzyme by ATP, so that when fATP] >> Kt the rate of ATP synthesis is proportional to l/[ATP] (Lehninger, 197.5, p. 198). The resulting rate of ATP synthesis (solid line in top panel) lags the hydrolytic rate by a time governed by r, the time constant for ATP turnover in the cell: eventually, the synthetic rate does catch up. The lower panel shows the e$ects of this process on extracellular acidtjication rate assuming that ATP is generated purely by glycolysis (solid line) or by glutamine oxidation (long dashed line: multiplied by nine compared to glycolysis). Glucose oxidation behaves similarly to glutamine oxidation. The short dashed line represents the time course for all carbon sources tfATP hydrolysis and synthesis are tightly coupled. Mathematical details: let c(t) be the instantaneous ATP concentration, initially c = co The rate of ATP synthesis then is (c&)[cdc(t)]. The rate ofATP hydrolysis was taken to be c&fort < 0 and (c&) (I +O.Jf(t))for t > 0, where f(t) = t3/[(0.2zp + t3]: f(t) modulates smoothly between 0 and 1. The resulting differential equation, dc/dt = (cd solved z)[c& - (I + 0.3f(t))], was numerically. Acidification rates were obtained according to Table I and section 4.2.1, neglecting the buffeting effects of cytoplasm. All quantities are plotted relative to their steady-state values for t < 0.

et al., 1978) and 75-120 s for Ehrlich ascites tumor cells (Skog etal., 1982). Measurements of ATP flux in MDCK cells by Lynch & Balaban (1987) suggest an intermediate time of about 30 s for these epithelial cells of kidney origin.

Biosensors & Bioelectronics

Another source of transients may involve changes in pHi_ Above we mentioned the observation that changes in pHi regulate metabolism in response to the activation of some receptors. Here we discuss not these metabolic effects, but the fate of the protons that are used to change the pHi itself. Although changes in pHi could arise from several sources, it appears that a major mechanism is alteration of the activity of the Na+/H+ exchange protein (e.g., Busa, 1986). If cytoplasmic alkalinization is achieved by an increased rate of proton excretion, how does this increase compare with basal rates of extracellular acidification? A simple calculation is useful. Internal pH is commonly observed to change by tenths of a unit on a time scale of minutes (Busa & Nuccitelli, 1984). In other words, rates of change of pHi on the order of 10e3 s-’ to lo-* s-’ are observed over minutes. Say that a cell has a volume of 2 pl and metabolically produces H+ at a steady rate in cytoplasm that has a buffer capacity of 30 mM/pH-’ (Roos & Boron (1981) cite measurements of buffer capacities ranging from about 15 to 80 mM, though Bountra et al. (1990) report that the upper values may be artifactual). This corresponds to proton excretion rates of z 10’ s-l and should be detectable if basal rates (also = 10’ s-r) are detectable on the minute time scale. These considerations are made more concrete in the more detailed model calculations displayed in Fig. 5. There the activity of the Na+/H+ exchanger is smoothly increased, leading to an increase in pHi that lags the exchanger activation due to intracellular buffering. The previously discussed dependence of glycolytic rate on pHi ensures that the steady-state acidification rate rises. The transient phase has two contributions: the modulation of glycolytic rate and, as described in the previous paragraph, the ejection of protons stored in the intracellular buffer as pHi rises. The result can be a transient overshoot of the extracellular acidification rate, as shown in Fig. 5. Note that the primary cause of this overshoot is independent of the rate of energy metabolism. Extracellular acidification rates, particularly transients, are manifestly sensitive to processes beyond energy metabolism. Two types of experiments support the contention that changes in pHi should be observable with the microphysiometer. First, Rice et al. (1991) used the microphysiometer to detect acidification transients associated with the

Biosensors based on the energy metabolism of living cells

o.“v-----l

0.9 d -I

0

1

?

3

Time, t/r

Fig. 5. Schematic modelfor the eflects of changes in intracellular pH on extracellular acidification rate. In this model, the rate of excretion ofprotons is a linearly decreasing function ofpH,, and the rate ofproduction ofprotons (e.g. by glycolysis) is a linearly increasing function of pH,. The pH at which the excretion and production rates are equal is a set-point pR and this pH is smoothly increased by 0.3 pH units by changing the parameters of the excretion-rate-vs.pH line beginning at t = 0; see the dashed curve in the top panel. This is intended to mimic the activation of the Na+/ H+ antiporter. ThepH, lags the instantaneous set-point pH but eventually reaches the new (higher) steady state. The time constant rfor the lag depends on the cytoplasmic buffer capacity, the cell volume, and the rates ofproton production and excretion: experimentally, time constants on the order of a minute are commonly observed. In the lower panel the increase in proton production (as pH, rises) is shown as a dashed line. The rate of proton excretion, which also includes the protons that had been stored in the cytoplasmic bufler. isshown as a solid line. Note that this modelgives an overshoot of acidtjication rate similar to that commonly observed in the microphysiometer when receptors are activated. Mathematical details: let y(t) be the instantaneous pH, and ySp(t) be the instantaneous set-point pH, (where production and excretion rates balance), both relative to some initial value prior to receptor stimulation. We chose ySp(t) = Ofor t < 0 and y+,,(t) = 0.3 f(t)for t > 0. with f(t) deJned as in Fig, 4. This behavior results from the underlying linearized model thatfor t > 0 the relative proton production and excretion rates are, respectively, (I + y)/r and [I + 0.6 f(t) - y]/r. where the numerical coeficients have been chosen for rough physical reasonableness. The pH, then obeys the dtferential equation dy/dt = -/y(t) y,(t)]/c This equation was solved numerically, and the valuesfor y(t) were used with the linearized production and excretion equations to generate the curves in the lower panel. All quantities are plotted relative to their steady-state values for t < 0. 269

J C. Owicki and J U? Parce

Biosensors & Bioelectronics

REFERENCES

acid-loading of cells exposed to weak acids such as pyruvate, caused by the selective permeability of the cell membrane to the protonated form of the acid. Second, H. G. Wada (personal communication) has shown that a cellular response to the cytokine granulocyte-macrophage colony-stimulating factor is in part due to activation of the Na+/H+ exchanger. He did so by observing the cytokine response with the microphysiometer when the principal extracellular cation was choline rather than Na+ and when the Na+/H+ exchanger had been inhibited by amiloride.

ships among platelet responses: studies on the burst in proton liberation, lactate production, and oxygen uptake during platelet aggregation and Ca*+ secretion. Blood, 57, 956-66. Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K & Watson, J. D. (1989).Molecular Biology of the Cell, 2nd. edn. Garland, New York. Andersen, 0. S. (1989).Elementary aspects of acid-base permeation and pH regulation.Ann. NYAcad. Sci.,

5. CONCLUSIONS

Balaban, R. S. & Bader, J. P. (1984). Studies on the relationship between glycolysis and (Na+ + K+> ATPase in cultured cells. Biochim. Biophys. Acta,

Akkerman, J. W. N. & Holmsen, H. (1981).Interrelation-

574, 333-53.

Balaban, R S. (1990). Regulation of oxidative phosphorylation in the mammalian cell. Am. .I. Physiol., 258, C377-89.

The foregoing discussion demonstrates that many disparate cellular processes can influence the extracellular acidification rate. This has three important consequences. First, it requires carefully controlled experiments to discover the mechanisms of acidification in a given system. These controls can be obtained with auxiliary analytical methods or by observing the effects of biochemical and genetic manipulations on acidification rate. Second, a wide variety of changes in the physiological state of living cells should produce changes in extracellular acidification rate. Third, biology plays a larger role here than it does in the engineering and development of more conventional biosensor systems. An instrument that can measure this rate conveniently with sufficient precision and time resolution, while allowing precise control of cell culture conditions, should be useful for applications ranging from general cell biology to in vitro toxicology and applications of ligandreceptor interactions such as drug discovery. At its present state of development, the silicon microphysiometer substantially fulfills these requirements.

ACKNOWLEDGMENTS We are pleased to thank the following colleagues for useful advice and discussions: C. Bountra, M. Deeg, J. Forte, N. Goldberg, J. Libby, L. Mandel, D. Miller, P. Rice, G. Sigal and G. Wada. Partial support from DARPA and CRDEC, AR0 Contract DAAL03-86-C-0009. 270

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Baron, R. (1989). Polarity and membrane transport in osteoclasts. Connect. Tissue Res., 20, 109-20. Borregaard, N., Schwartz, J. H. & Tauber, k I. (1984). Proton secretion by stimulated neutrophils. J. Clin. Invest., 74, 455-9.

Bountra, C., Powell, T. & Vaughan-Jones, R D. (1990). Comparison of intracellular pH transients in single ventricular myocytes and isolated ventricular muscle of guinea-pig. J. Physiol. (London), 424, 343-65. Bousse, L., Parce. J. W., Owicki. J. C. & Miller, K M.

(1990). Silicon micromachining in the fabrication of biosensors using living cells. Tech. Dig. IEEE Solid State Sensor and Actuator Workshop, 173-6. Bruner, L. H., Kain, D. J., Roberts, D. A. & Parker, R D. (1991a). Evaluation of seven in-vitro alternatives for ocular safety testing. Fund. Appl. Tax., 17,136-49. Bruner, L. H., Miller, K M., Owicki. J. C., Parce, J. W. & Muir, V. C. (1991 b). Testing ocular irritancy in vitro with the silicon microphysiometer. Toxicol. In Vitro, 5, 277-84. Busa, W. B. (1986).The proton as an integrating effector in metabolic activation. Cum. Top. Membrane Transport. 26, 291-305. Busa, W. B. & Nuccitelli, R (1984). Metabolic regulation via intracellular pH.Am. J. Physiol., 246, R409-38. Butcher, F. R, McBride, P. A. & Rudich, L. (1976). Choline@ regulation of cyclic nucleotide levels, amylase release, and K+ efflux from rat parotid glands. Mol. Cell. Endocrinol., 5, 243-54. Chung, F.-Z., Wang, C.-D., Potter, P. C., Venter, J. C. & Fraser, C. M. (1988). Site-directed mutagenesis and continuous expression of human /3-adrenergic receptors. J. Biol. Chem., 263, 4052-5.

Clegg, J. S. & Gordon, E. P. (1985). Respiratory metabolism of L-929 cells at different water contents andvo1umes.J. Cell Physiol, 124,299-304. Dawson, R. M. C., Elliott, D. C., Elliott, W. H. &Jones,

Biosensors & Bioelectronics

Biosensors based on the energy metabolism of living cells

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