Chemotactic Responses of Escherichia coli to Small - Europe PMC

Chemotactic Responses of Escherichia coli to Small - Europe PMC

1706 Biophysical Journal Volume 76 March 1999 1706 –1719 Chemotactic Responses of Escherichia coli to Small Jumps of Photoreleased L-Aspartate Ra...

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Biophysical Journal

Volume 76

March 1999

1706 –1719

Chemotactic Responses of Escherichia coli to Small Jumps of Photoreleased L-Aspartate Ravi Jasuja,* Jinsoo Keyoung,* Gordon P. Reid,# David R. Trentham,# and Shahid Khan* *Department of Physiology and Biophysics, Albert Einstein College of Medicine, Bronx, New York 10461 USA, and #National Institute for Medical Research, Mill Hill, London NW7 1AA, England

ABSTRACT Computer-assisted motion analysis coupled to flash photolysis of caged chemoeffectors provides a means for time-resolved analysis of bacterial chemotaxis. Escherichia coli taxis toward the amino acid attractant L-aspartate is mediated by the Tar receptor. The physiology of this response, as well as Tar structure and biochemistry, has been studied extensively. The b-2,6-dinitrobenzyl ester of L-aspartic acid and the 1-(2-nitrophenyl)ethyl ether of 8-hydroxypyrene-1,3,6-tris-sulfonic acid were synthesized. These compounds liberated L-aspartate and the fluorophore 8-hydroxypyrene 1,3,6-tris-sulfonic acid (pyranine) upon irradiation with near-UV light. Photorelease of the fluorophore was used to define the amplitude and temporal stability of the aspartate jumps employed in chemotaxis experiments. The dependence of chemotactic adaptation times on aspartate concentration, determined in mixing experiments, was best fit by two Tar aspartate-binding sites. Signal processing (excitation) times, amplitudes, and adaptive recovery of responses elicited by aspartate jumps producing less than 20% change in receptor occupancy were characterized in photorelease assays. Aspartate concentration jumps in the nanomolar range elicited measurable responses. The response threshold and sensitivity of swimming bacteria matched those of bacteria tethered to glass by a single flagellum. Stimuli of similar magnitude, delivered either by rapid mixing or photorelease, evoked responses of similar strength, as assessed by recovery time measurements. These times remained proportional to change in receptor occupancy close to threshold, irrespective of prior occupancy. Motor excitation responses decayed exponentially with time. Rates of excitation responses near threshold ranged from 2 to 7 s21. These values are consistent with control of excitation signaling by decay of phosphorylated pools of the response regulator protein, CheY. Excitation response rates increased slightly with stimulus size up to values limited by the instrumentation; the most rapid was measured to be 16 6 3 (SE) s21. This increase may reflect simultaneous activation of CheY dephosphorylation, together with inhibition of its phosphorylation.

INTRODUCTION Strong chemotactic responses are evoked by acidic amino acids in a wide variety of sensory systems ranging from bacteria to humans. Chemotaxis of the bacterium Escherichia coli toward amino acid chemoattractants provides a well-understood example of a single-cell sensory response (Berg, 1983; Blair, 1995; Stock and Surette, 1996; Appleby et al., 1996; Falke et al., 1997; Bray, 1998). The motility of these bacteria consists of an alternating pattern of swimming runs and tumbles. During runs, the six or so flagella per cell form a counterclockwise (CCW) rotating bundle that provides thrust. Reversal to clockwise (CW) rotation of an undetermined number of flagella breaks up the bundle, causing tumbling events that randomize cell orientation. Chemotactic migration is accomplished by increasing swimming runs up spatial, attractant gradients (Berg and Brown, 1972). The smooth-swim response to L-aspartate is mediated by the Tar receptor, one of a family of transmembrane methylaccepting chemotaxis proteins (MCPs) that mediate responses to diverse stimuli. Insights into Tar structure and its modulation by aspartate binding have been obtained by Received for publication 12 August 1998 and in final form 18 November 1998. Address reprint requests to Dr. Shahid Khan, Department of Physiology and Biophysics, Albert Einstein College of Medicine, Bronx, NY 10461. Tel.: 718-430-4046; Fax: 718-430-8819; E-mail: [email protected] © 1999 by the Biophysical Society 0006-3495/99/03/1706/14 $2.00

x-ray crystallography (Milburn et al., 1991), NMR (Danielson et al., 1994), and mutagenesis (Falke and Koshland, 1987; Cochran and Kim, 1996; Gardina and Manson, 1996). Motile responses to changes in extracellular aspartate concentration are effected by an intracellular signaling cascade composed of the chemotaxis proteins CheA, CheW, and CheY. Tar and other MCPs form complexes with the linker CheW and the histidine kinase CheA (Gegner et al., 1992). CheA, as part of such receptor complexes, phosphorylates CheY, forming CheY-P. CheY regulates motor rotation, binding to the motor when phosphorylated and promoting its CW rotation. Aspartate binding to Tar inhibits CheA-to-CheY phosphotransfer, lowering the concentration of CheY-P. This promotes CCW rotation and hence smooth swimming. The chemotactic excitation is transient. Simultaneous initiation of other processes leads to adaptation back to the prestimulus state. Glutamic acid residues in the Tar cytoplasmic domain are continually methylated and demethylated by the transferase CheR and the esterase CheB respectively. CheB has a regulatory domain, homologous to CheY, which, when phosphorylated by CheA, inhibits its catalytic domain. The resulting net increase in methylation leads to adaptation. CheY-P dephosphorylation is accelerated by CheZ protein. Modulation of CheZ’s aggregation state may play a role in adaptation (Blat and Eisenbach, 1994; Eisenbach, 1996) and/or signal amplification (Wang and Matsumura, 1996).

Jasuja et al.

Chemotactic signaling in E. coli

Chemotactic excitation and subsequent adaptation may be separated in time by monitoring responses to rapid changes in chemoeffector concentration. This has been accomplished for free-swimming bacteria by mixing (Macnab and Koshland, 1972) and flash photolysis of photosensitive “caged” chemoeffectors (Khan et al., 1993). Concentration changes effected by flow-cell exchange (Berg and Tedesco, 1975) and iontophoresis (Segall et al., 1982) were used to study responses of bacteria tethered by a single flagellum to glass coverslips. Adaptation times of responses to the large step stimuli studied by such methods ranged from tens of seconds to minutes and were proportional to fractional changes in receptor occupancy (Spudich and Koshland, 1975). When chemoeffector concentrations were changed enzymatically (Brown and Berg, 1974) or ramped (Block et al., 1983), such that both excitation and adaptation contributed to the response, the response amplitude was proportional to the rate of change of receptor occupancy. Only flash photorelease and iontophoresis were sufficiently rapid for direct measurement of the subsecond signal processing or excitation times. Excitation kinetics measured by flash photorelease and iontophoresis were similar for responses that saturated (i.e., produced completely smoothswimming populations). However, there were discrepancies in response sensitivity, as assessed from the amplitudes of subsaturation responses. A response latency, evident in the iontophoretic measurements, was not observed upon flash photorelease of serine or protons (Khan et al., 1993, 1995). The present study was undertaken to provide a more detailed description of chemotactic response kinetics and resolve these discrepancies. MATERIALS AND METHODS


5.2. The pH was adjusted to 4.4 with aqueous HCl, and a further 0.3 mmol diazoethane was added in 1.5 ml CHCl3. The shaking was continued for a further 5 h, at the end of which 8% of the HPTS remained. The reaction mixture was then partitioned between CHCl3 and water at pH 7.8. The aqueous phase was loaded onto a DEAE cellulose column (2.5 cm diameter 3 50 cm) and eluted with a 4-liter gradient of 0.01–1.01 M TEAB (triethylammonium bicarbonate) at pH 7.4 and 5°C. After some minor impurities, HPTS eluted as a broad peak (centered at 0.68 M TEAB) fairly well resolved from caged HPTS (centered at 0.8 M TEAB) and an unknown contaminant at .0.9 M TEAB. HPTS was distinguished from caged HPTS because only the former absorbs at 454 nm (e 2.01 3 104 M21 s21 at pH . 8). This absorbance was used to identify and discard aliquots of caged HPTS containing HPTS. The unknown contaminant had an absorption band at 373 nm. It was identified as present in the eluate when the ratio of absorption at 403 nm to that at 373 nm was less than 1.23. For caged HPTS this ratio was 1.28. Tubes containing pure caged HPTS were pooled and evaporated to dryness in vacuo several times after addition of methanol to the dried material to remove TEAB. The tris-triethylammonium salt of caged HPTS was obtained in 70% overall yield from HPTS and stored in the dark at 220°C in aqueous solution. Quantification was based on e 1.85 3 104 M21 s21 at lmax 403 nm in aqueous solution near neutrality (pH 6 – 8). Qp, the product quantum yield of caged HPTS on photolysis was measured by comparison with that of the P3-1-(2-nitrophenyl)ethyl ester of ATP (caged ATP), using the approach described by Walker et al. (1989). In a typical experiment a cuvette containing 0.10 mM caged HPTS, 0.10 mM caged ATP, 2 mM dithiothreitol, and 50 mM ammonium phosphate buffer at pH 7.0 and 22°C was illuminated with a xenon arc lamp for various times through a UG11 320 (6 40)-nm bandpass filter. Samples of the solution were analyzed by fluorescence and/or absorption spectroscopy at pH 10 to detect formation of HPTS (e 2.01 3 104 M21 s21 at 454 nm, a wavelength at which caged HPTS does not absorb) and by high-performance liquid chromatography (HPLC) (Whatman Partisil SAX) to measure formation of ATP and decay of caged HPTS and caged ATP (HPTS was retained on the column), using aqueous 0.54 M (NH4)H2PO4/(NH4)2HPO4 at pH 5.5 and acetonitrile (10% by volume) as the eluting solvent. The rate of photolysis of caged HPTS was measured by recording the absorption or fluorescence changes associated with the formation of HPTS after flash photolysis (20-ms pulse of a Candela dye laser at 320 nm) (Walker et al., 1988).

Caged compounds New caged compounds, caged HPTS, the O-1-(2-nitrophenyl)ethyl ether of 8-hydroxypyrene-1,3,6-tris-sulfonic acid, caged L-aspartate (Scheme 1), and the P3-2,6-dinitrobenzyl ester of ATP (2,6-dinitrobenzyl-caged ATP) were synthesized for these studies. Caged HPTS is a nonfluorescent compound that on photolysis releases fluorescent HPTS. This fluorescence was used to measure the extent of caged HPTS photolysis and thus to calibrate the photolysis apparatus. The intermediate leading to HPTS at a rate with constant kHPTS is an aci-nitro compound (Barth et al., 1997, and references therein). Caged L-aspartate also photolyzes via an aci-nitro intermediate that has a broad absorption band. Its exponential decay with rate constant kasp probably determines the rate of L-aspartate formation (Corrie and Trentham, 1993). The nitrosoketone by-products react with thiols and were rendered biologically inert by including dithiothreitol (DTT) in the solution, as done previously for other caged chemoeffectors (Khan et al., 1993).

Synthesis and properties of caged HPTS The approach used to synthesize caged HPTS has been described by Walker et al. (1989). The hydrazone of 2-nitroacetophenone (311 mg (1.76 mmol)) was quantitatively oxidized to 1-(2-nitrophenyl)diazoethane. Ddiazoethane (1.45 mmol) in 7.3 ml CHCl3 was added to 1 mmol of the trisodium salt of HPTS (pyranine; Lancaster Synthesis, Morecombe, England) dissolved in 5 ml H2O at pH 4.86. The mixture was shaken vigorously in a stoppered flask for 4 h at 21°C. At this time, absorption spectra indicated that 22% of the HPTS remained, and the pH had risen to

Synthesis and properties of caged aspartate (b-2,6dinitrobenzyl ester of L-aspartic acid) N-t-BOC-a-t-butyl L-aspartate (Sigma Chemical Co., St. Louis, MO) (0.5 mmol) was treated with 1 mmol KF and 0.5 mmol 2,6-dinitrobenzyl bromide under reflux in dry acetone for 4 days. The reaction was followed by thin-layer chromatography on K6F plates, monitoring the appearance of a UV-absorbing spot. The t-BOC ester was purified by flash chromatography. After removal of solvent the product was treated with 5 ml trifluoroacetic acid for 1 h at 22°C and rotary evaporated to dryness, followed by 4 evaporations with methanol. Caged aspartate was obtained in 68% yield, stored in the dark at 220°C in methanol, and assayed from e 12,000 M21 cm21 at lmax 230 nm. Amino acid analysis of material, redissolved in water at pH 3 after evaporation of methanol, contained 1.4% aspartic acid. 2,6-Dinitrobenzyl esters are prone to spontaneous hydrolysis in aqueous solution at a rate proportional to hydroxide ion concentration. The rate of caged aspartate hydrolysis was 1.1 3 1024 min21 at 22°C and pH 7.0. During an experiment it was kept in solution on ice wrapped in foil until required, so as to minimize hydrolysis. Qp of caged aspartate was also measured by comparison with caged ATP. First the 2,6-dinitrobenzyl chromophore was compared with that of the 1-(2-nitrophenyl)ethyl group, the photolabile moiety of caged ATP and caged HPTS, in the wavelength range transparent to the UG11 bandpass filter. The spectra are almost identical from 335 nm to 360 nm, but from 300 nm to 335 nm the 2,6-nitrobenzyl group only absorbs ;75% as effectively on average. This difference was ignored because the illumina-


Biophysical Journal

Volume 76

March 1999

Scheme 1.

tion for photolysis used in the microscope assays was similar to that used for spectroscopic measurements. The photolysis protocol was as used above for caged HPTS, except that analysis was by HPLC, using a Merck RP8 reverse-phase column with aqueous 5 mM KH2PO4/K2HPO4 at pH 5.6 and methanol (10% by volume) as the eluting solvent. Caged ATP, caged aspartate, ATP, and the byproduct of caged aspartate photolysis were monitored at 254 nm to follow the photolysis. The photolysis of caged HPTS and caged aspartate was also compared directly because the aspartate photoreleased in the biological assays was measured from the fluorescence of HPTS generated by caged HPTS photolysis. Similar spectroscopic and HPLC analytical methods (RP8 reverse phase column) were used as when comparing caged HPTS and caged aspartate with caged ATP. The rate of photolysis of caged L-aspartate was inferred from the rate of decay of the aci-nitro intermediate formed after flash photolysis (Walker et al., 1988), using the same apparatus as for caged HPTS.

Synthesis and properties of the P3-2,6-dinitrobenzyl ester of ATP (2,6-dinitrobenzyl-caged ATP) 2,6-Dinitrophenyldiazomethane was prepared from the parent aldehyde according to the methods of Walker et al. (1989). 2,6-Dinitrobenzyl-caged

ATP was synthesized from ATP and the diazomethane in a mixed CHCl3/ water solvent at pH 4 and then isolated in 15% overall yield, using DEAE anion exchange chromatography and preparative reverse-phase HPLC. 2,6-Dinitrobenzyl-caged ATP had Qp 5 0.6 when measured against caged ATP as standard (see comment above on Qp measurement of 2,6-dinitrobenzyl compounds). On flash photolysis ATP formed at 10 s21 at 22°C, 0.2 M ionic strength, and pH 7, as measured by the rate of decay of the aci-nitro intermediate.

Growth and preparation of bacteria E. coli strains RP437 (wild type for motility and chemotaxis) and RP2361 (Tar deletion mutant) were obtained from Dr. J. S. Parkinson. Overnight cultures were prepared from bacteria that were motility selected on tryptone soft agar (0.35% agar). Inocula (1/100) of the overnight cultures were grown at 35°C in tryptone broth (plus 20 mg/ml streptomycin). The bacteria were harvested in late exponential phase (OD600 nm ' 0.5 cm21). They were washed three times with buffer A (20 mM Na2HPO4/KH2PO4, pH 7.0 6 0.2, 10 mM potassium chloride, 0.1 mM EDTA, 5 mM lithium

Jasuja et al.

Chemotactic signaling in E. coli

lactate, 125 mM methionine). For caged aspartate experiments, buffer A contained, in addition, 5 mM DTT. For tethered-cell experiments, the bacteria were sheared (27-gauge needles) after the first wash, then washed two more times. The washed bacteria were incubated with flagellar antibody-coated circular (22 mm) coverslips for ;30 min. The coverslips were mounted on a laminar flow cell (Berg and Block, 1984). Untethered bacteria were subsequently washed out by buffer A.

Mixing experiments A custom-made mixing apparatus was utilized. This consisted of two reservoirs, one filled with buffer A containing aspartate and the other with buffer A containing bacteria, which fed into a 10-ml mixing chamber. The mixed solution was pulled via valve (PS3; Pharmacia Biotech, Uppsala, Sweden) through the laminar flow cell observation chamber with a peristaltic pump (Gilson minipuls II) at a rate of 0.03 ml/s. The mixing ratio and wash-in/wash-out times of the mixed suspensions were determined by mixing HPTS solutions of known concentration with buffer A. Mixing was accomplished within 0.1 s. The transit time between mixing and exchange into the chamber was 7 6 1 s at the flow rate used. Higher (more than fivefold) flow rates impaired the motility of the bacteria, probably because of breakage of flagella. Flow was stopped for motility measurements by simultaneously turning off the pump and closing the valve controlling entry to the flow cell.

Photorelease assays Photolysis of caged HPTS was used to measure the energy of the photolyzing flash and, thus, estimate the amounts of caged chemoeffectors, here aspartate, photolyzed in behavioral assays. The fluorescence of known concentrations of HPTS (Molecular Probes, Eugene, OR) in aqueous 0.1 M phosphate buffer at pH 7.0 was used to construct a calibration curve for determination of amounts of caged HPTS photolyzed in the microscope. A 450 6 15-nm exciter/510-nm dichroic/.520-nm barrier filter cassette was used for excitation and visualization of HPTS fluorescence. The intensity was recorded using a photodiode (Thorlabs model 201/579-7227) connected to a current-to-voltage amplifier (Oriel 70710), digital readout (Oriel 70701), and a chart recorder (Pharmacia REC101). Alternatively, video records were made and fluorescence intensities obtained upon digitization (Khan et al., 1993). From the fluorescence released on photolysis of caged HPTS, the amount of aspartate released was calculated. Two modes with different geometries were used to effect photolysis of caged chemoeffectors in microscope behavioral assays (Khan et al., 1993). In one mode, light from a Gert-Rapp flash lamp was transmitted through a UG11 filter with infrared reflective coating and directed onto the sample at an angle of 40 6 2° via a liquid light guide that terminated in a planoconvex lens. The collimated beam from the lens illuminated all or most of the sample. In the other mode, shuttered episcopic Kohler illumination from a continuous mercury arc lamp (Nikon), also directed through the UG11 filter, effected photolysis over a local area comparable to the microscopic field of view as seen through the eyepieces. Distinction between diffusion out of the video field of view and photobleaching of released HPTS was achieved by varying the viscosity of the solution with glycerol. Viscosity of HPTS/glycerol buffers was measured using a Cannon-Ubbelohde (size 100) viscometer. The bacteria were imaged using a long working distance objective (323 magnification, 0.5 numerical aperture) when photorelease was effected by direct side illumination from a flash lamp and by a fluorite phase-contrast objective (403 magnification, 0.85 numerical aperture) when epiillumination effected photorelease. The transmissivity of the fluorite objective (;65% at 360 nm) decreased at lower wavelengths. For swimming-cell assays, samples of ;5 ml volume were observed in a bridged coverslip chamber through a quartz coverslip positioned on two glass (0.17 mm) coverslips. The quartz coverslips were cleaned in fuming


nitric acid before use. This treatment reduced the number of bacteria stuck to the coverslip surface. For tethered-cell assays, the flow cell was used. After bacteria were introduced, buffer with caged aspartate was introduced and 10 min was allowed for equilibration before flash photorelease. The free aspartate contamination in the caged aspartate was also determined using a bioassay based on CheB mutants. CheB mutants have abnormally long adaptive recovery times (Clarke and Koshland, 1979). Recovery times of Salmonella typhimurium SL4041 CheB mutant strain upon mixing with known aspartate concentrations were measured. A calibration curve constructed from these measurements was used to read off the aspartate concentration from the responses observed when the mutant bacteria were mixed with samples of caged aspartate. The contamination level was 1.5%, in correspondence with values determined after synthesis, implying that degradation during subsequent handling was negligible. Addition of D-aspartate, a nonmetabolizable analog, was used to increase prestimulus receptor occupancy, which was always nonzero because of L-aspartate contamination of the caged sample. The gain (g) provided a measure of response strength. This, as defined previously (Khan et al., 1993), was D(motor bias)/DRocc; where D(motor bias), the change in {CCW/(CW 1 CCW)}rotation bias, was estimated from the change in the rate of change of direction (Drcd), given D(motor bias) 5 2(0.0012)Drcd (figure 2 of Khan et al., 1993).

Motion analysis Video records were digitized off-line (VP320 digitizer, ExpertVision version 1.4 software; Motion Analysis, Santa Rosa, CA) and analyzed as previously described (Khan et al., 1993, 1995). A zoom lens coupled to the video camera was used to adjust the final magnification such that one pixel in the digitized image spanned 0.5 mm. For each concentration change, the aggregate response of .1000 bacteria was measured: a single flash sequence typically captured responses of 50 –100 bacteria. Swimming cell records were batch-processed at 30 and 15 frames/s for measurement of excitation response (t1/2) and adaptation recovery (tr) times, respectively. These were the times required for a half-maximum response and recovery back to prestimulus behavior, respectively (Khan et al., 1995). Excitation response data were expressed as the response rate, kex, where kex 5 ln 2/t1/2. Because at 15 frames/s, linear speed (spd operator) as well as the frame-to-frame angular speed (rcd operator) is sensitive to changes in swim-tumble behavior (Sager et al., 1988; Khan et al., 1993), the rcd/spd ratio, as well as the rcd alone, was used to measure recovery times. Under our experimental conditions, unstimulated wild-type bacteria typically had a population rcd between 750 and 850°/s. This corresponded to a {CCW/ (CW 1 CCW)} motor rotation bias of 0.7 and 0.6, respectively. Excitation responses were fitted by single exponentials, using nonlinear least-squares curve-fitting routines available in SIGMAPLOT (Jandel Scientific) software. Nonlinear least-squares procedures were also used to infer KD’s by fitting models to the recovery time data obtained in mixing experiments. For estimation of peak response amplitudes and recovery times, successive 10-frame (i.e., 0.33 s) windows were averaged. The number of windows used for determination of peak response amplitude depended on the duration of the postexcitatory period before initiation of adaptive recovery. For tethered-cell experiments, the unavoidable presence of stuck cells precluded batch processing of the video data. Single tethered bacteria were boxed out and digitized individually at 60 frames/s. One pixel spanned 0.3 mm in the digitized images. Analysis was limited to bacteria with rotation rates under 20 Hz. Avel and ngdr operators were used to compute the angular velocity and monitor regularity of rotation. The avel is the first derivative of the direction of travel with respect to time. The ngdr is the ratio of the distance from the first point in the path to the given point over the actual path distance from the first point to the given point. The rotation sense, given by the sign of the avel, was subsequently extracted. Individual cell path files were then merged to yield the time dependence of the population change in motor rotation bias.


Biophysical Journal

RESULTS Spectroscopy and photochemistry of caged compounds The extent of caged aspartate photolysis relative to that of caged HPTS when the compounds were exposed to near-UV light in a cuvette was used to estimate the aspartate concentration jumps delivered in microscope photorelease assays. Caged HPTS has a pH-independent absorption spectrum over the 4 –10 pH range and is not fluorescent (Fig. 1). The excitation spectra of HPTS overlap with its absorption spectra in the 350 –500-nm range. The invariance of the emission spectra in the 4 –10 pH range is due to the fact that the pKa of the excited state is ;1 (Parker, 1968). The extents of caged aspartate and caged HPTS photolysis relative to that of caged ATP were 33% and 20%, respectively. In separate experiments, direct comparison of caged aspartate and caged HPTS photolysis confirmed the expected ratio of 1.6 (33%/20%) within a 620% error range. Qp, the number of product molecules divided by the number of photons absorbed at a particular wavelength or, as here, wavelength range was 0.63 for caged ATP (Walker et al., 1988, 1989). Thus the Qp’s for caged aspartate and caged HPTS are 0.21 and 0.13, respectively. The estimated Qp for caged HPTS does not take into account absorption of near-UV photons by the pyrene ring system as well as by the 1-(2-nitrophenyl)ethyl chromophore. However, this does not affect the calibration of the extent of caged aspartate photolysis using caged HPTS. Proton release studies on photolysis of 2,6-dinitrobenzyl acetate (Nerbonne, 1986) suggested Op values for 2,6-dinotrobenzyl carboxylate esters are significantly greater than those of 1-(2-nitrophenyl) ethyl esters. By contrast, Op valves for 2,6-dinitrobenzyl

Volume 76

March 1999

and 1-(2-nitrophenyl) ethyl caged ATPs were within 10% of one another. The rates of photolysis of caged HPTS and caged aspartate were measured from the appearance of the absorption or fluorescence of HPTS and from the decay of the aci-nitro intermediate, respectively (Fig. 2). The rates of formation of HPTS and aspartate were 550 s21 and 630 s21, respectively, at pH 7.0, 0.1 M ionic strength, and 22°C. They were approximately proportional to proton concentration over the 5–9 pH range. Stimulus characteristics The two microscop