Combining Ultrarapid Mixing with Photochemical Oxidation to Probe

Department of Physics and Astronomy, Michigan State University, East Lansing, Michigan 48824, United States. Anal. Chem. , 2013, 85 (10), pp 4920–49...
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Combining Ultrarapid Mixing with Photochemical Oxidation to Probe Protein Folding Ling Wu and Lisa J. Lapidus* Department of Physics and Astronomy, Michigan State University, East Lansing, Michigan 48824, United States S Supporting Information *

ABSTRACT: We demonstrate a new method to study protein folding by combining fast photochemical oxidation of proteins (FPOP) with ultrarapid microfluidic mixing to observe kinetics on the microsecond time scale. Folding proteins pass through a focused UV laser beam, creating OH radicals that label the select protein side chains and are analyzed with mass spectrometry. As a proof of principle, we demonstrate this method with hen egg lysozyme that shows at least two kinetic phases before 1 ms, which are compared with those observed by Trp fluorescence. This method provides another, complementary probe of the early, complex steps of protein folding.

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FPOP and temperature jump on the submillisecond time scale.8,9 This experiment used a “pump/probe” two-laser setup in which one laser provided a temperature jump within 10 ns and the other generated OH radicals with a variable delay. However, the lifetime of the OH radical in their protocol is about 1 μs, which limited the observation to microsecond time scales, not nanoseconds typically accessible by T-jump. Furthermore, most proteins can only be unfolded by T-jump rather than folded from a fully unfolded state. In this work, we couple FPOP with a rapid microfluidic mixing device in which a fully unfolded protein starts folding after mixing within 8 μs and is labeled by OH radicals along the folding process. With this technique, the kinetics of protein solvent accessibility during folding can be followed with microsecond time resolution.

he complexity of protein folding has motivated development of many techniques to observe the process over a wide range of time scales, including laser temperature jump, pressure jump, stopped flow mixing, and ultrarapid continuous mixing; there have been parallel developments for probes to observe the process, including hydrogen-exchange NMR, circular dichroism, intrinsic tryptophan fluorescence, and fluorescence resonance energy transfer (FRET). Often combining these prompt and probe techniques presents considerable technical challenges, usually due to the sensitivity of the probe or the time scale of the prompt. This has been a particular challenge for stopped-flow mixing in which typically single-shot data are collected with millisecond resolution. In contrast, continuous flow mixing allows observation at submillisecond resolution but can use significant amounts of sample.1,2 Microfluidic mixing overcomes these hurdles by scaling mixer dimensions down to the laminar flow regime such that sample consumption is small and the mixing time is low. To observe folding, typically the fastest microfluidic mixers have been coupled with fluorescence because it is easy to detect in a small volume. However, a fluorescent probe is usually localized to a specific position in the protein, and ideally we would prefer a more general probe to monitor the folding behavior of the entire molecule. Fast photochemical oxidation of proteins (FPOP)3 is a powerful chemical footprinting technique to study the protein folding process.4 One application is to use a laser to initiate photolysis of hydrogen peroxide to produce OH radicals which label exposed amino acid residues.3 After detection of modified residues (including Met, Cys, Trp, Phe, Tyr, His, Pro, Leu, and Ile) with mass spectrometry, solvent accessibility of the protein can be characterized. Recently, Stocks and Konermann characterized structural changes of cytochrome c, holo-myoglobin, and S100A11 during folding by combining stopped-flow mixing with FPOP on the millisecond time scale.5−7 Gross and co-workers have also measured protein folding dynamics of barstar by combining © 2013 American Chemical Society

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EXPERIMENTAL SECTION Microfluidic Mixer. The basic design of this mixer was first described by Knight et al.10 and was optimized by Hertzog et al.11 (see Figure 1a). This design takes advantage of small channel dimensions to keep the dynamics in the laminar flow regime, even for very high velocities, resulting in mixing times of as low as 8 μs (see Figure 1c, gray points and blue line)12,13 (some data were collected with lower flow rates and longer mixing times to achieve a longer observation time). The fluid dynamics in the chip are simulated with Comsol Multiphysics (Comsol, Stockholm, Sweden). The channels were etched in 500-μm-thick fused silica wafers using reactive ion etching with polysilicon as a mask. Inlet and outlet holes were drilled with a diamond-tipped drill. The channels were then first prebonded to a 170-μm-thick fused silica wafer after a reverse RCA cleaning and then fused together at 1100 °C. The mixer is Received: November 20, 2012 Accepted: April 17, 2013 Published: April 17, 2013 4920

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focused to a 1 μm spot by a 0.5 NA UV objective (OFR 40x266, Newton, NJ) inside the mixer. For a linear flow speed of 1 m/s, the 1 μm spot results in a maximum time resolution of 1 μs. Fluorescence intensity is collected by the same objective and sent through a dichroic mirror (Chroma 300dclp, Brattleboro, VT) to a photon counter (Hamamatsu H742140, Hamamatsu City, Japan). To collect fluorescence spectra, the beam is diverted to a spectrograph (Jobin Yvon MicroHR, Horiba, Ltd., Edison, NJ) and CCD camera (iDus 420A-BU, Andor Technology, Belfast, UK). The mixer manifold is mounted on a three-axis piezoelectric scanner (Mad City Laboratories Nano-LP100, Madison, WI), which scans the chip over the objective 100 μm in each direction and a motorized microscope stage (Semprex Corp, Campbell, CA) that moves the chip by 80 μm down the channel. Hen-egg lysozyme was purchased from Sigma (St. Louis, MO) and used without further purification. Lysozyme (500 μM) was unfolded by dissolving in 6 M guanidine HCl, and folding was initiated by mixing with 100 mM potassium phosphate buffer (pH 7). A typical experiment begins with a scan of the exit channel imaged by the photon counter to locate the jet of fluorescent protein. The chip is typically scanned 10 μm across the exit channel and 500 μm down the linear section of the exit channel (see Figure 1b). This corresponds to ∼500 μs of folding time at an initial flow rate of 1 m/s without substantial diffusion of the protein out of the jet. Alternatively, long time courses can be obtained by slowing the flow rate to as low as 0.2 m/s. The overall intensity as a function of time can be obtained from a scan by averaging the fluorescence intensity of the jet in ∼1 μm2 regions, the size of the excitation beam. Fast Photochemical Oxidation. The setup of FPOP is similar to the fluorescence measurement. Lysozyme (500 μM) with 6 M GdnCl flows through the center channel, and folding is initiated by mixing with potassium phosphate buffer from the side channel. In addition, 15 mM hydrogen peroxide was added to the side channels to provide hydroxyl radicals upon the laserinduced photochemical reaction, following the protocol in Hambly et al.3 H2O2 is only added to the side channels because the smaller center channel dimensions would clog with oxygen gas bubbles produced from H2O2. To reduce the OH radical lifetime from at least 100 μs3 to ∼1 μs, 20 mM glutamine was added in both center and side channels as a scavenger for the hydroxyl radical.16 Figures S1 and S2 (Supporting Information) show the mass spectra of folded and unfolded lysozyme in FPOP with and without glutamine and show that the level of oxidation depends inversely on the concentration of the scavenger. The 258 nm laser with ∼5 mW power is focused onto a 1 μm region inside the exit channel of mixer. The actual size of the jet is calculated to be ∼100 nm,11 so all protein flowing past the laser beam is exposed to light. In that 1 μm region, hydrogen peroxide is photolyzed to produce hydroxyl radicals, and exposed amino acid residues of protein are oxidatively modified by reaction with OH radicals. The laser sits at the same spot on the jet for a period of minutes to accumulate labeled protein from one point in experimental time. Different laser powers and flow rates were tested to ensure saturated photolysis of hydrogen peroxide (Figures S3 and S4, Supporting Information). A 100 μL solution can be collected in 20 min at a flow rate of 1 m/s. This volume is sufficient for a good signal from mass spectrometry, but more would be needed if the protein concentration is lower. The laser is then moved to another position, and another sample is collected. Each sample is collected in an Eppendorf tube

Figure 1. Observation of lysozyme folding in a microfluidic mixer by Trp fluorescence. (a) Electron micrograph of fused silica mixer centered on the mixing region. The protein in denaturant enters from the top left corner and is met by flows from either side channel containing the mixing buffer. The protein is hydrodynamically focused to a narrow jet, the denaturant molecules are diluted 100-fold, and all the streams proceed down the observation channel (bottom right corner) at a continuous rate. The observation channel is 10 μm wide, 10 μm deep, and 500 μm long. (b) Contour plot of intensity of Trp fluorescence within the channel. The protein flows from top to bottom and is met by mixing buffer at ∼80 μm. (c) (Black points) Total Trp fluorescence intensity of lysozyme, relative to a nonfolding control experiment, as a function of time. The red line is a two-exponential fit to the data. (Gray points) Total Trp fluorescence of Nacetyltryptophan amide after mixing into 400 mM KI, a diffusionlimited fluorescence quencher, relative to a control experiment with 0 mM KI, as a function of time. The blue line is the concentration of denaturant as a function of time calculated using COMSOL.

mounted on a manifold which contains solution reservoirs for each channel in the chip. The flow rate of each channel is controlled by air pressure above the reservoir using computercontrolled pressure transducers (Marsh Bellofram Type 2000, Newell, WV). At the fastest flow rates reported, the sample consumption is ∼4 μL/h of the protein and 400 μL/h of folding buffer.14 Fluorescence Detection. The UV fluorescence of a folding protein is monitored with a specially designed confocal microscope.15 An argon-ion laser (Lexel 95-SHG, Cambridge Laser, Fremont, CA) at 257 nm enters an inverted microscope (Olympus IX51, Melville, NY) as a collimated beam and is 4921

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containing 20 μL of 100 nM catalase and 70 mM methionine to remove excess H2O2 after FPOP. The sample was left at room temperature for 10 min before storing at 5 °C to allow the complete decomposition of hydrogen peroxide by catalase. The samples were concentrated and desalted with a C18 ZipTip prior to mass spectrometry analysis. Mass Spectrometry. A Waters (Milford, MA) Quattro Premier mass spectrometer was coupled to a Acquity HPLC system. A Waters Symmetry Beta Basic CN column (10 × 1 mm, 5 μm particle size) was used at room temperature. The injection volume was 10−20 μL, and the HPLC flow rate was 0.1 mL/min, achieved by using the gradient from 2% acetonitrile (98% water) to 75% acetonitrile over 12 min and then back to 2% acetonitrile followed by a 3 min reequilibration step. Formic acid (0.1%) was added into the water mobile phase to ensure solubility. Mass spectra were acquired by using electrospray ionization in the positive ion mode. The capillary voltage, extractor voltage, and cone voltage were set at 3.17 kV, 5 V, and 25 V, respectively. The flow rates of the cone gas and desolvation gas were 30 and 600 L/h, respectively. The source temperature and desolvation temperature were 120 and 350 °C, respectively. Data were acquired with MassLynx 4.1 and processed for calibration and for quantification of the analytes with QuanLynx software.

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RESULTS Measuring FPOP during Folding. The chip, shown in Figure 1a, is mounted on a confocal microscope for positioning the laser beam within the channel. To position the beam accurately, the mixing region is first scanned using UV fluorescence detection. Figure 1b shows a contour plot of the intensity in the mixing region measured by the photon counter. After the protein jet is located using fluorescence, the laser beam is moved to a specific point along the jet and sample is collected in an Eppendorf tube until 100 μL is collected, which takes about 20 min at a flow rate of 1 m/s. At this flow rate, the protein jet, which also contains 15 mM H2O2 and 20 mM glutamine, crosses the ∼1 μm diameter beam in ∼1 μs. OH radicals are created by the UV light at 258 nm and are quenched in about ∼1 μs by the 20 mM glutamine.3 Higher concentrations of glutamine could shorten this lifetime further, but this lifetime is commensurate with the time resolution of the mixer. Thus, each sample represents a ∼1 μs slice in the folding process. The beam is then moved to another point along the jet, and sample is collected in another tube. After concentrating and desalting the sample, it is analyzed using liquid chromatography and mass spectrometry. Figure 2a−d shows four representative spectra before, during, and after folding of hen-egg lysozyme. Each normalized spectrum shows multiple peaks corresponding to the mass of the unmodified protein (14306 MW) plus multiples of 16 MW, the mass increment when an H atom in the protein is replaced with the OH. The spectrum of the unfolded protein is dominated by the peak at protein+2 radicals, indicating that the chain is fairly solvent exposed. The spectrum at 20 μs after mixing shows a significant increase in the unlabeled protein, while the spectrum at 480 μs shows a shift back to higher mass. Finally the spectrum of the folded protein again shows a spectrum dominated by the unmodified protein. The integrated intensity of each of the first three peaks (summed over 16 mass units around the peak) is plotted in Figure 2e. Each shows a significant jump within the mixing region, a decay in the peak corresponding to unmodified protein (and a corresponding rise

Figure 2. (a−d) Mass spectra of FPOP-labeled lysozyme at various times before, during, and after folding as marked. The amplitude is normalized to be 1 over the entire spectrum. (e) The total intensity over 16 mass units of the first three mass peaks vs time. The black lines are single exponential fits with decay/rise times given in the main text.

in the singly and doubly modified protein) over 500 μs and then a rise in the unmodified protein (and a drop in the singly and doubly modified protein) after 500 μs and before the protein folds. The exponential decay/rise times for the unlabeled and 1 OH peaks are 165 ± 16 μs and 176 ± 38 μs (see Figure 2e for fits). The first measured time for the unlabeled protein peak indicates an additional faster decay on the time scale of mixing. The doubly modified protein peak appears to show a much longer rise time that is not fully resolved within the 500 μs window. Measuring Fluorescence during Folding. The observed folding of lysozyme using FPOP can be compared with observations made of fluorescence from lysozyme’s six tryptophans. The contour plot in Figure 1b measures the total fluorescence intensity over time as the protein moves down the exit channel. After scanning vertically within the exit channel in 100 μm increments, the intensity of emitted light along the jet is extracted and plotted vs time after mixing, where the time is calculated from the jet position and the exit channel 4922

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flow rate. To correct for a large decrease in fluorescence in the mixing region due to formation of the protein jet, a control experiment is performed in which the protein in 6 M GdnHCl is mixed with buffer containing 6 M GdnHCl as well, and the fluorescence is observed. This trace is divided point-by-point into the folding trace. Figure 1c shows the relative fluorescence intensity of the six tryptophan residues in lysozyme as a function of time. The decay curves at different flow rates agree well (except within the first ∼20 μs because the mixing times are different). The data after 12 μs are fit to two exponential decays with lifetimes 36 ± 2 μs and 180 ± 15 μs. The second phase is commensurate with the rates measured by FPOP. There is also an apparent rise in signal within the mixing time relative to the unfolded state. An alternative probe of folding is the global analysis of the fluorescence spectral changes with time. Figure 3a shows the spectra collected over time. The large peak at early time is the unfolded protein before the mixing region. After mixing, the signal decreases significantly (due to formation of the jet) and shifts to lower wavelengths (due to decreased solvent exposure of Trp). Analysis of all spectra with single value decomposition (svd) yields three significant components (all other components are approximately 10-fold smaller than the third component and show no significant change with time). The first component (blue) in Figure 3b is the average spectrum over all time which shows similar kinetics (Figure 3c) to the total intensity measurement (Figure 1c), but because it is dominated by the large decrease in signal during mixing, the data are not fit numerically. The second (green) and third (red) components are the time-dependent deviations from the average spectrum due to the change in solvent exposure of the tryptophans which results in a blue-shift of the fluorescence spectrum over time. That there are two spectral shift components indicates that not all of the six tryptophans change conformation uniformly. The time dependence of the second component mostly changes within the mixing time, although there is a small exponential rise on the 100 μs time scale. The third component shows a rapid rise within the mixing time, a decay on the order of the mixing time (9.3 ± 0.4 μs) and a further decay with a lifetime of 264 ± 5 μs (see Figure 3c for fit). The rise in signal represents a blue-shift (low solvent exposure) in the spectrum and a decay represents a redshift (high solvent exposure). This suggests at least some of the tryptophans rapidly become less solvent exposed and then gradually resolvate over the first 1 ms of folding, similar to the changes observed by FPOP.

Figure 3. (a) Time-dependent Trp fluorescence spectra measured before, during, and after mixing. (b) Three most significant spectral components from svd analysis. The singular values are 1.19 × 106 (blue), 2.14 × 105 (green), and 7.93 × 103 (red). (c) Time dependence of the three most significant svd components measured for a flow rate of 0.5 m/s. (d) Time dependence of the third svd component (red in panels b and c) measured at three different flow rates (1, 0.5, and 0.25 m/s) and combined. The red line is a fit of the combined data to two exponential decays.

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DISCUSSION Measuring FPOP on the entire folding protein chain is in itself an excellent complement to conventional probes such as fluorescence or CD, and in the future the use of protease digestion and amino acid level mass spectrometry should give even more specific structural information.9 However, interpreting such data as indicative of a well-defined state should be done with caution, given the complexity of kinetics observed here. It is likely that at any time point there will be multiple populations of labeled sites, reflecting multiple pathways at the earliest stages of folding. This was observed by Chen et al., measuring barstar folding after T-jump, in which measurements of each labeled site yielded large uncertainties, although measurement error cannot be ruled out.9 Figures S1 and S2 show that under high OH radical quenching conditions that give microsecond time resolution, the number of modified sites

is reduced 2- to 3-fold compared to no quenching. That is because not all residues label at the same rate and efficiency (and some do not label at all). Thus, a lack of labeling at a particular site does not preclude that it is solvent exposed. Interpreting the fluorescence intensity, spectral shift and FPOP data together suggest significant complexity in the folding kinetics of lysozyme. All three probes show a rapid change with the mixing and then a turnaround in the signal over 0.5−1 ms. The rates of change of total fluorescence intensity and two FPOP peaks are in agreement but disagree with the fluorescence spectral shift and the third FPOP peak. 4923

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ACKNOWLEDGMENTS We thank Michael Gross and Olgicia Bakajin for many helpful discussions, Stephen DeCamp for fabrication of the mixers, and the RTSF Mass Spectrometry Core at MSU for the support of collecting and processing of all mass data. This work is supported by the National Science Foundation NSF DBI0754570.

The rate of the third FPOP peak may be in agreement with the rate of the third svd component of the fluorescence spectra, but the time range of the FPOP data precludes quantitative comparison. Additionally there is another, 40 μs kinetic phase in the total fluorescence intensity. Altogether, this data suggests at least four kinetic phases before 1 ms. An earlier study has already revealed some complexity on the submillisecond time scale. Bachmann et al. showed that an intermediate state with more helical structure than the native state formed within the 1.2 ms dead time of their stopped-flow mixer,17 followed by a turnaround in the CD signal. The FPOP measurements reveal some surprising results regarding the solvent accessibility of the protein in the early stages of folding. During the mixing time, the number of labeling sites decreases, suggesting significant collapse with dilution of the denaturant, an effect that has been seen with several proteins previously by Trp spectral shift or fluorescence resonance energy transfer.12,15,18,19 However, the slower phase on the 100 μs time scale indicates that some residues are actually becoming more solvent exposed as seen by the increase in the number of modification sites. It is not clear that the protein is actually less compact after 500 μs; SAXS experiments by Bachmann et al. show a continuous decrease in the radius of gyration on the millisecond time scale. It is known that water is a poor solvent for both hydrophobic residues20 and the protein backbone,21,22 so these phases suggest that the unfolded chain is rapidly reconfigured during the change of solvent to sequester both from water, but this conformation is perhaps not productive for folding, so the structure loosens slightly, before finding a metastable intermediate structure. Recent experimental and computational work on the denatured state of the protein NTL9 shows substantial intramolecular interactions even as the unfolded ensemble retained qualities of a completely random coil. This implies that hydrophobic burial can occur without substantial compaction of the entire chain.23 Finally, there is at least one final folding step after 1 ms in which the native structure packs properly, causing solvent exposure to decrease. The final folding time has been measured by multiple probes to be ∼200 ms, but the kinetics of the third mass peak (2 OH labels) suggest there is another phase on the millisecond time scale which could be commensurate with the intermediate identified by Bachmann et al.17 In summary, we have shown that time-resolved FPOP after ultrarapid mixing reveals new information about protein folding. In the case of lysozyme, we find the surprising result that after large-scale collapse due to the change in solvent, the protein becomes somewhat more solvent exposed, possibly to facilitate the formation of correct native contacts. Together with kinetic data from Trp fluorescence, there is strong evidence of multiple kinetic phases and perhaps multiple pathways in the earliest stages of folding.

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ASSOCIATED CONTENT

S Supporting Information *

Additional information as noted in the text. This material is available free of charge via the Internet at http://pubs.acs.org.

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Notes

The authors declare no competing financial interest. 4924

dx.doi.org/10.1021/ac3033646 | Anal. Chem. 2013, 85, 4920−4924