Article pubs.acs.org/ac
Ultrasensitive and Specific Measurement of Protease Activity Using Functionalized Photonic Crystals Bakul Gupta,† Kelly Mai,‡ Stuart B. Lowe,† Denis Wakefield,‡ Nick Di Girolamo,‡ Katharina Gaus,§ Peter J. Reece,∥ and J. Justin Gooding*,† †
School of Chemistry, The Australian Centre for NanoMedicine and ARC Centre of Excellence in Convergent Bio-Nano Science and Technology, UNSW Australia, Sydney, New South Wales 2052, Australia ‡ School of Medical Sciences, UNSW Australia, Sydney, New South Wales 2052, Australia § EMBL Australia Node in Single Molecule Science, School of Medical Sciences and ARC Centre of Excellence in Advanced Molecular Imaging, UNSW Australia, Sydney, New South Wales 2052, Australia ∥ School of Physics, UNSW Australia, Sydney, New South Wales 2052, Australia S Supporting Information *
ABSTRACT: Herein is presented a microsensor technology as a diagnostic tool for detecting specific matrix metalloproteinases (MMPs) at very low concentrations. MMP-2 and MMP-9 are detected using label free porous silicon (PSi) photonic crystals that have been made selective for a given MMP by filling the nanopores with synthetic polymeric substrates containing a peptide sequence for that MMP. Proteolytic cleavage of the peptide sequence results in a shift in wavelength of the main peak in the reflectivity spectrum of the PSi device, which is dependent on the amount of MMP present. The ability to detect picogram amounts of MMP-2 and MMP-9 released by primary retinal pigment epithelial (RPE) cells and iris pigment epithelial (IPE) cells stimulated with lipopolysaccharide (LPS) is demonstrated. It was found that both cell types secrete higher amounts of MMP-2 than MMP-9 in their stimulated state, with RPE cells producing higher amounts of MMPs than IPE cells. The microsensor performance was compared to conventional protease detection systems, including gelatin zymography and enzyme linked immunosorbent assay (ELISA). It was found that the PSi microsensors were more sensitive than gelatin zymography; PSi microsensors detected the presence of both MMP-2 and MMP-9 while zymography could only detect MMP-2. The MMP-2 and MMP-9 quantification correlated well with the ELISA. This new method of detecting protease activity shows superior performance to conventional protease assays and has the potential for translation to high-throughput multiplexed analysis.
T
Along with the synthesis and secretion of a number of proinflammatory cytokines, corneal epithelial cells produce and release a variety of MMPs, such as gelatinases (MMP-2 and -9), collagenases (MMP-1 and -13), and stromelysins (MMP-3, -10, and -11).6,7 Therefore, accurate assessment of these enzymes, once released from their cell of origin, is important for understanding the mechanism of disease development. A variety of techniques including fluorometric assays8,9 and colorimetric assays10 have been developed to measure protease activity. Gel zymography has been the predominant method employed.11−13 However, a limitation of this technique is that it does not discriminate between different MMPs because these enzymes have overlapping substrate specificities.14 Other challenges with this technique include the staining and destaining of acrylamide gels which can often lead to loss of
he expression of protease enzymes, or their endogenous inhibitors that regulate protease activity, can be readily determined at the transcript level with whole genome microarray chips.1 Determining which cells are expressing them, and in what amounts, is often critical to understanding their functional roles. Therefore, it is important to recognize the quantities in which individual cells and cell population produce proteases for the development of effective therapies. One such class of proteases, the matrix metalloproteinase (MMP), is an integral part of the immune system. Macrophages release MMPs, and macrophage cell activities are closely related to acute and chronic inflammation.2,3 Therefore, detecting MMPs released by these cells can provide information about the status of immune activation. Another example where detection and quantification of MMPs can be critical is the dry eye syndrome. It has been proposed that it is the synthesis and secretion of pro-inflammatory cytokines and MMPs by the corneal epithelium, which contributes to the development of ocular surface inflammation in patients with this condition.4,5 © 2015 American Chemical Society
Received: July 6, 2015 Accepted: August 27, 2015 Published: August 27, 2015 9946
DOI: 10.1021/acs.analchem.5b02529 Anal. Chem. 2015, 87, 9946−9953
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peptide surface chemistry35 and extends the technology to be able to quantify the levels of MMP-2 and MMP-9 released from primary cells for the first time. The methodology involves the modification of PSi surfaces with 1,8-nonadyine, followed by the copper-catalyzed “click” cycloaddition of the distal alkyne with an antifouling polymer, poly(ethylene glycol acrylate-statacrylic acid) containing an azide moiety. This step was then followed by the addition of the peptide sequences, VPLSLYSGK (specific for MMP-2) and SGKGPRQITAK (specific for MMP-9), which were further linked to another polymeric species, poly(hydroxyethyl acrylate-stat-N-hydroxysuccinimide ester acrylate), that served as a sacrificial polymer that leaves the pores after the MMPs cleave the peptide linkage. The purpose of this paper is to demonstrate the application of this new technology as an assay method to profile MMP levels from primary cell culture by quantifying the amounts of MMP-2 and MMP-9 released by cells. Specifically, primary retinal pigment epithelial (RPE) and iris pigment epithelial (IPE) cells were investigated because of their link to ocular diseases such as dry eye syndrome. Work done by Nagai et al.36 and Lan et al.37 showed that RPE and IPE cells release MMPs on stimulation with the bacterial endotoxin lipopolysaccharide (LPS). We show here that MMP cleavage of specific peptide sequences resulted in an optical shift in the reflectivity spectra of the photonic microsensors to shorter wavelengths (blue shift). The performance of PSi microsensor based sensors was compared to the conventional methods of gelating zymography and ELISA.
sensitivity or bleaching of the bands making them unsuitable for quantification.15 An alternative technique that is employed in conjunction with zymography is enzyme linked immunosorbent assay (ELISA). Although ELISAs are very sensitive and can detect and quantify proteases at low levels, they do not provide information on the activity of the enzyme and therefore cannot differentiate between active and inactive forms within a sample. Porous silicon (PSi) is a promising material for its use in cell assays due to its biocompatibility,16−18 tunable architecture, and versatility for optical biosensing.19−30 It is made from pure single crystalline silicon by electrochemical etching. It can be fabricated to reflect light at well-defined wavelengths. The position of these reflectivity peaks is sensitive to any change in the refractive index of the bulk PSi material. Its common application in biosensing is to place affinity molecules such as antibodies into the pores and monitor the change in optical reflectivity when the antigen selectively binds to the pore walls of the PSi. However, more recently PSi sensors have been shown to be able to monitor enzyme activity.31−33 PSi has been employed as an MMP specific sensor in a number of configurations. For example, Orosco et al.23 fabricated a PSi rugate filter that was made hydrophobic in the interior by methylation of the micropores. The hydrophobic protein zein was adsorbed onto the top surface of the porous film, and the underlying pores remained air-filled. Incubation with the protease pepsin led to zein digestion and subsequent infiltration of the small peptide fragments and water into the vacant pores. The pore loading caused an increase in the refractive index of the PSi layer, which in turn, induced a measurable red shift to the reflectance band. With this biosensor, protease concentrations as low as 7 μM could be detected. Furthermore, with protease concentrations higher than 14 μM, the changes to the porous matrix were so profound that they could be observed by the naked eye. In another study, Martin et al.34 used antibody functionalized PSi resonant microcavities to detect the presence of MMP-8. This was achieved by observing the shift in the resonance dip cavity of the PSi structure. The device achieved MMP-8 detection down to 1.5 nM. Both these sensors demonstrate the operation in pure buffers rather than in complex biological fluids. Responding to the challenge of monitoring in biological fluids, recent work by Krismastuti et al.33 demonstrated the use of the PSi microcavity to detect MMP-1 from wound fluids. The PSi microcavity was functionalized using a fluorogenic MMP peptide substrate featuring both a fluorophore and a quencher. Once the active MMPs recognize and cleave the peptide sequence of the substrate, it produced an immobilized peptide fragment carrying the fluorophore yielding a fluorescence signal. The reported limit of detection with this fluorescence based PSi biosensor in wound fluid samples was 1.5 × 10−15 M. A previous study by Kilian et al.19 demonstrated the use of PSi rugate filter to detect the gelatinase MMPs secreted by viable human macrophages. The innovation in this device was to fill the entire pore space with gelatin as a substrate that responded to both MMP-2 and MMP-9. The impressive aspect of this device was the lowest detected concentration of gelatinases at 0.37 pM, an unprecedented low detection limit. Advancing this work, we recently developed PSi rugate filters where the pores were filled with a synthetic polymer system that can be cross-linked with any given peptide sequence. In this way, specific peptides sequences allow the discrimination of the levels of MMP-2 from MMP-9 and vice versa.35 The work presented herein uses the same polymer−
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EXPERIMENTAL PROCEDURES Materials. All chemicals were of analytical grade or higher and purchased from Sigma-Aldrich (Sydney, Australia) unless otherwise stated. Lipopolysaccharide (LPS, from E. coli 0111:B4) was purchased from Sigma-Aldrich; active MMP-2 and MMP-9 were purchased from Calbiochem and used as received. Chemicals used in surface modification procedures were of high purity (≥99%). 1,8-Nonadiyne (Alfa Aesar, 97%) was redistilled from sodium borohydride (Sigma-Aldrich, 99+ %) under reduced pressure (79 °C, 8−9 Torr), collected over activated molecular sieves (Fluka, 3 Å pore diameter, 10−20 mesh beads, dehydrated with indicator), and stored under a dry argon atmosphere prior to use. Milli-Q water (>18 MΩ·cm) was used to prepare solutions and for chemical reactions. Dichloromethane and ethanol for surface cleaning, chemical reactions, and purification procedures were redistilled prior to use. 50% hydrofluoric acid used for porous silicon etching was obtained from Ajax Finechem (Taren Point, Australia). Prime grade single-side polished silicon wafers, 100-oriented (⟨100⟩ ± 0.05°), p-type (boron), 500−550 μm thick, 0.001−0.0015 Ω cm resistivity as provided by suppliers, were obtained from SILTRONIX (Archamps, France). Porous Silicon Rugate Filter Synthesis. Porous silicon (PSi) rugate filter samples were prepared in a custom-made electrochemical cell made from Teflon with 35% hydrofluoric acid (HF)/absolute ethanol solution (1:1, v/v) as described previously.20,35 Current densities varying between 169.6 and 229.7 mA·cm−2) were applied periodically to result in a porous structure with 60 periods with their porosity varying from 57% to 60%. Note: this 3% porosity difference does not yield a large difference in the pore diameter of the two layers as observed in Figure S1a. For lift-off samples, a high current pulse (320 mA, 100 ms) was employed subsequently in a 15% (v/v) ethanolic solution 9947
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streptomycin and maintained at 37 °C in an atmosphere of 5% CO2. For stimulation assays, the primary cells were seeded at a density of 3 × 104 cells/mL in 24-well plates (NUNC, Denmark) and used for experimentation once they reached confluence. The modified PSi microsensors were added to the confluent cells in a 24-well plate. The cells were then stimulated with different doses of LPS (0, 1, 2, 5, and 10 μg·mL−1) and cell supernatants applied in PSi protease assays, zymography, and ELISAs. For protease assays, the sensors’ reflectivity was measured when the microsensors were incubated in supernatants collected from cells or in vitro, that is, when the sensors were present in the cell culture media along with the cells. For zymography and ELISA, culture supernatants (∼250 μL) were collected at different time points (0, 2, 4, 6, 10, 16, 24, and 48 h) into microfuge tubes and stored at −80 °C until assayed. Gelatin Zymography. Gelatin substrate zymography was modified from the procedure published by Di Girolamo et al.41 Porcine type A gelatin (Sigma) was added to a standard 10% acrylamide resolving gel mixture at a final concentration of 1 mg·mL−1. Enzymatic activities were identified as clear zones (lytic bands) in a blue-stained background. MMP Detection Using ELISA. The human MMP-2 and MMP-9 ELISA kits were purchased from Sigma-Aldrich and used within a week of their delivery. For MMP-2, the culture supernatants were diluted 5−25-fold, whereas for MMP-9 it was 500−1000-fold. The dilutions were carried out using the 1× diluent buffer provided in the kit. Supernatant from nonstimulated cells (i.e., with no LPS) was used as positive controls (to determine the basal level of proteases secreted) whereas culture medium alone was used as a negative control. The assay was performed in accordance with the manufacturer’s instructions. Sample dilutions were run in duplicate. Error bars present in ELISA graphed results indicate standard deviation of these measurements. Optical Reflectivity Measurements. Optical reflectivity spectra of PSi microsensors were measured at normal incidence using a custom built optical arrangement plus an inverted Leica DM IL LED microscope (Leica Microsystems Pty Ltd., USA). The USB2000+ miniature fiber optic spectrometer (Ocean Optics Inc., USA) was coupled to a multimode fiber with the collecting facet mounted in the conjugate image plane of the microscope. The microscope was equipped with an N PLAN L 40×/0.55 air objective, an epi-illumination bright field slider, and a ProgRes CFscan camera (JENOPTIK Laser, Optik, System GmbH, Germany). Reflectivity spectra were also recorded by LabVIEW and processed using Prism 5.0 (GraphPad Software, Inc., USA). Scanning Electron Microscopy. Surface and cross-sectional scanning electron micrographs (SEMs) of the PSi structures were recorded prior to lifting off and sanitation using a FEI Nova NanoSEM 230 field emission scanning electron microscope (SEM). The microscope was operated with an extraction voltage of 5 kV, had a spot size of 3 nm, and employed an in-lens secondary electron (SE) detection system. For the SEM of PSi microsensors, the ultrasonicated microsensors were dried onto a silicon 0.5 cm × 0.5 cm wafer attached to a steel stub and were imaged using FEI Nova NanoSEM 230. Optical Microscopy. The optical micrographs of PSi microsensors were obtained by placing PSi microsensors on a glass slide and using an Olympus BX53 optical microscope. The microscope was equipped with Olympus XC50 for imaging these PSi microsensors.
of HF, to make the porous structure stand free from the substrate but with the circumference of the PSi remaining attached to the substrate. The PSi microsensors were prepared by sonicating this electrochemically lifted-off PSi film off the silicon substrate. Surface Modification of PSi Rugate Filters. Formation of monolayers by hydrosilylation of an alkyne to form a stable Si−C bond was achieved using a thermal reaction in a neat alkyne solution of 1,8-nonadiyne following a previously reported procedure.38 The alkyne modified PSi structures were further modified with a polymer that resists nonspecific adsorption of proteins (antifouling polymer) via “click” chemistry. In a typical “click” procedure, to a reaction vial containing the alkyne-functionalized PSi surface were added (i) the azide polymer poly(ethylene glycol acrylate-stat-acrylic acid) (50 mg in 1 mL of ethanol), (ii) copper(II) sulfate pentahydrate (1 mol % relative to the azide), (iii) sodium ascorbate (4 mg·mL−1 in 1 mL of Milli-Q water), and (iv) N,N,N′,N′-tetramethylethane-1,2-diamine (TMEDA, 20 μL). Reactions were conducted at room temperature and stopped after 15 to 16 h. The antifouling polymer modified samples were further modified to activate the carboxylic acid moiety to a reactive succinimide ester moiety. 1-Ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC, 0.50 mmol) and N-hydroxysuccinimide (NHS, 1.05 mmol) were dissolved in Milli-Q water (5 mL), and the PSi was added to this solution for 3 h. The modified PSi samples were then transferred to a microfuge tube containing the peptide sequence of choice. The two peptide sequences used for MMP-2 and MMP-9 were VPLSLYSGK and SGKGPRQITAK, respectively. Peptide immobilization was achieved by incubating the activated sample with 1 mg·mL−1 peptide in 1× PBS for 3−4 h. The last coupling step was the addition of the sacrificial polymer, poly(hydroxyethyl acrylate-stat-N-hydroxysuccinimide ester acrylate). Typically, to a reaction vial, 80−100 mg of the synthesized polymer was added to 1 mL of Milli-Q water. To this reaction vial, the peptide-modified samples were added. The reaction was allowed to take place for a period of 16 h yielding the polymer−peptide−polymer surface, which was rinsed with copious amounts of Milli-Q water. Protease Digestion Assays. For all the samples, where the protease solution was added to the microsensors in a Petri dish, the protease assays were performed in wet conditions; that is, the measurements were conducted in phosphate buffered saline (PBS), pH 7.1−7.4. Approximately 150 μL of protease solution was required to cover all the microsensors in a Petri dish. For the experiments where proteases were released by the cells, the sensors and cells were typically incubated in 2 mL of serum free cell culture medium at 37 °C. The reflectivity measurements were performed at various time points, and for the measurement, the Petri dishes containing the sensors were taken out of the 37 °C incubator. Cell Culture and Stimulation. Donor matched IPE and RPE cells were isolated from human eyes (3 donors) and cultured as described previously.39 Donor human eyes were obtained from Lions NSW Eye Bank, Sydney, Australia. The level of proteins released by epithelial cells from different patients is variable.40 Therefore, in order to remove variability and maintain consistency between various studies, both the RPE and IPE cells were collected from the same human donor. Briefly, the primary cells were cultured in epithelial cell medium (ScienCell Research Laboratories, San Diego, CA) supplemented with 2% FBS, 100 U·mL−1 penicillin, and 100 mg·mL−1 9948
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Determination of Kinetic Parameters Using PSi Microsensors. The kinetics of the responses of the PSi microsensors to the two enzymes of interest, the gelatinases MMP-2 and MMP-9, were characterized. The changes observed in the optical shifts, such as those depicted in Figure S3, were expressed as the optical response of the sensor where optical response was defined as
RESULTS AND DISCUSSION The optical structures used for the MMP assays described herein are called rugate filters. Rugate filters are characterized by having a sinusoidal variation in porosity across the structure, which results in them only reflecting a narrow band light with other wavelengths being absorbed by the structure.42 The PSi rugate structures in this study had 60 layers and were formed by electrochemical etching of p-type silicon. The cross section of the pores shows they are columnar but have some branching (see Figure S1a). The average pore diameter calculated from the top view scanning electron micrographs (see Figure S1b) was 34 ± 10 nm. To turn PSi rugate filters into MMP microsensors requires filling the pore space of the PSi with an enzyme sensitive substrate, which in this case is an enzyme degradable peptide. This is because the position of the reflectance band is not only dependent on the fabrication variable such as pore size but also dependent on the average refractive index of the material.42 Typically, organic matter is of higher refractive index than water. Hence, when enzymes enter the pores of the PSi, they degrade the enzyme sensitive peptide and cause it to leave the pores. As the water that replaces the polymer in the pores is of lower refractive index, the reflectance peak shifts to shorter wavelengths (see abstract graphic), referred to as a blue shift. To turn the as fabricated PSi photonic crystals embedded in a silicon wafer into microparticles, the chip containing the PSi is subjected to a high current pulse, which lifts the PSi off the silicon.43,44 The lift-off sample is then subjected to ultrasonication to give PSi particles shown in the SEM image in Figure S2a. The fact that the porous structure of the PSi is not destroyed in this fabrication process can be seen from the porosity still evident in Figure S2b. The PSi microparticles are then turned into PSi microsensors by first modifying them with surface chemistry that stabilizes the silicon from degradation in biological media (see Scheme S1). The surface chemistry involved reacting the molecule 1,8-nonadiyne with the PSi via thermal hydrosilylation followed by coupling a polymer system that possesses an enzyme degradable peptide cross-linker, to the surface chemistry in the pores of the PSi.35 The evidence that the PSi microsensors retain their optical properties during these different fabrication steps is clear from the red brown color of the final PSi sensors shown in the optical micrographs in Figure S2c,d. The ability of the PSi microsensors to respond to MMPs is shown in Figure S3. In a typical experiment, the PSi microsensors were incubated with MMP-2 or MMP-9 in PBS buffer for a period of 24 h. The MMP-2 or MMP-9 that enter the pores digests the peptide sequences of the polymer−peptide construct, and the degradation products leave the pores which changes the average refractive index of the photonic microsensors. This change in the average refractive index of the microsensors results in the shifting of the reflectivity peak toward shorter wavelengths (blue shift) in the reflectivity spectrum of the microsensor.35 Such changes are evident in Figure S3a where a blue shift in the reflectance band of the microsensor occurs when incubated in a solution containing MMP-2 for 24 h. The peak position of the reflectivity peak at 24 h was compared to the peak position of the reflectivity peak at 0 h to yield the magnitude of blue shift observed. In contrast, in the absence of MMP-2, that is when the PSi is simply exposed to buffer, a negligible blue shift (∼1 nm) was obtained (Figure S3b).
optical reponse(%) =
Δλprotease − Δλcontrol Δλpeptide + polymer
× 100 (1)
where Δλprotease was the blue shift obtained when sensors were present in the MMP-2 or MMP-9 solution; Δλ control corresponds to the blue shift observed when the sensors were present in PBS containing no enzyme, and Δλpeptide+polymer refers to the initial shift in the reflectivity spectrum when the PSi microsensor pores were filled with the peptide and polymer. The optical responses obtained when the MMP-2 and MMP9 enzyme specific peptides containing microsensors were incubated with different MMP-2 and MMP-9 enzyme concentrations are shown in Figures 1a and S4a. After 5 h of
Figure 1. (a) The optical response obtained when polymer−peptide modified PSi microsensors (n = 3) were incubated with different concentrations of MMP-2 enzyme (i, 1 nM; ii, 10 nM; iii, 50 nM; iv, 100 nM; v, 375 nM; vi, 750 nM; viii, 1.5 μM). The data was fitted to an exponential decay function for all different concentrations. (b) The curve obtained when the V0/[S] was plotted against the MMP-2 enzyme concentrations used. (c) The slope was calculated from the first 4 linear points from the data obtained in (b) to yield the kcat/KM parameter for the MMP-2 protease.
incubation with the proteases, no further change in optical response was observed. This was true for both the high concentrations (1.2 μM for MMP-9 and 1.5 μM for MMP-2) and low concentration of enzyme (1 nM), which means that after 5 h there was no more degradation of the peptide by the proteases. This was attributed to the loss of activity of the enzyme after 5 h, as distinct from all the peptide−polymer substrate being consumed, as the latter would result in a close to 100% optical response. The optical responses at low levels of protease enzyme demonstrates the applicability of the PSi microsensors for detecting different MMPs at low nanomolar concentrations. To study surface enzymatic reaction rates quantitatively, Kim et al. have examined the reaction of collagenase on peptide 9949
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donor eye were used. The cells were stimulated with 1 μg·mL−1 of a bacterial endotoxin LPS, and the supernatant was collected at 2, 6, and 24 h post LPS addition. The PSi sensors were then incubated with the collected cell supernatants with their reflectivity being measured at 24 h. Figure 2 displays the blue
monolayers45,46 and the reaction of protease on BSA monolayers.47 For the case of protease adsorption and reaction, they proposed a modified Michaelis−Menten model for the surface kinetics. Michaelis−Menten kinetics has also been shown to be applicable for PSi based protease sensors by Kilian et al.19 In the latter study, mass transport was not rate limiting and interfacial substrate concentration was far in excess of the amount of the enzyme because of the large volume of substrate these nanoporous materials possess.19 That is, mass transport of the enzyme to the substrate could be neglected and only enzyme kinetics needed to be considered. Therefore, in order to study the enzyme kinetics, the optical responses were fitted to an exponential decay function (Figures 1a and S4a). The slope of the optical response curves give the velocity, V0/[S], which is the rate at which the enzyme reacts with the substrate to form the enzyme−substrate complex. This velocity was plotted as a function of input MMP concentration (Figures 1b and S5b). Since the V0/[S] curve is linear only at low enzyme concentrations, only the concentrations within the linear range were used to calculate the kcat/KM value for the MMP (from the slope of the curve in Figures 1c and S4c). The calculated kcat/KM for MMP-2 was (6.5 ± 0.6) × 104 −1 M s−1. This value is in excellent agreement with the previously reported value of (6.1 ± 0.4) × 104 M−1 s−1 calculated using a conventional solution based assay.48 In this solution based procedure, the peptide cleavage was assayed by following the production of amine using fluorescamine. The amount of product was determined by using the signal from a given peptide digested to completion with a standard protease. The kcat/KM was determined directly from the initial rate at a single substrate concentration under conditions where KM ≫ [S].48 In a similar manner, the kcat/KM was calculated for the MMP9 specific peptide modified PSi microsensors (Figure S4) and the kcat/KM value obtained from this data was (1.62 ± 0.13) × 105 M−1 s−1. This calculated value was also in good agreement with the value of 1.88 × 105 M−1 s−1 determined directly from the initial rate at a single substrate concentration under conditions where KM ≫ [S] in a solution phase assay.49 The kcat/KM ratio obtained for MMP-9 was 2.5 times higher than the ratio obtained for MMP-2 suggesting a higher affinity of the enzyme for the peptide entrapped inside the PSi matrix. The good agreement between the kinetic parameters for MMP-2 and MMP-9 obtained using PSi rugate filters with the values obtained using a solution based assay shows that mass transport was not a limiting factor. This was attributed to the fact that the enzyme is in excess relative to the number of pores and that the enzyme diffuses rapidly through the polymers in the pore spaces. Agreement between these results and those in the literature offers support for the validity of this PSi optical biosensor method. Finally, the results presented in these calibration curves also present the broad range of concentrations these microsensors can respond to, which make them a potential tool to be used to determine enzyme activity in vitro. Detection of MMP-2 and MMP-9 Released by Primary Cells Using PSi. Detection of MMPs from ocular epithelial cells is critical for diagnosing diseases like dry eye syndrome50 and inflammatory diseases like uveitis.51−53 Therefore, the utility of the polymer−peptide modified PSi microsensors for profiling MMP enzymatic activity from ocular epithelial cells was evaluated. The levels of MMP-2 and MMP-9 released from primary cells were determined by collecting the supernatant from cells in culture. The RPE cells derived from a human
Figure 2. Blue shifts obtained after the 24 h incubation of (a) MMP-2 and (b) MMP-9 specific PSi microsensors (n = 5) with the cell supernatants. The supernatant was collected from LPS stimulated and nonstimulated RPE cells at different times of 2, 6, and 24 h.
shifts obtained after the 24 h incubation of PSi microsensors in RPE cell supernatants collected from RPE cells stimulated with and without LPS. The PSi microsensors used in Figure 2a were modified with MMP-2 specific peptide sequence whereas, in Figure 2b, the microsensors were made responsive to only MMP-9. In Figure 2a,b, it was observed that the magnitude of blue shift, obtained when the microsensors were incubated in the supernatant of cells stimulated with LPS, was higher when compared to the blue shift obtained from incubating microsensors in supernatants collected from RPE cells not stimulated with LPS. The relatively small blue shifts observed in the absence of LPS were attributed to the endogenous levels of MMP-2 and MMP-9 secreted by these cells as previously reported by Ahir et al.54 and Eichler et al.55 The magnitude of blue shift also increased with the increase in time after which the supernatant was collected from the cells. The higher magnitude of blue shift indicates higher amounts of MMP-2 and MMP-9 present in the supernatant collected from LPS stimulated cells. This was expected because LPS is known to upregulate the expression of MMP-2 and MMP-9 in RPE cells upon LPS stimulation. Gelatin zymography was used to compare the sensitivity of the potential use of PSi as a biosensor for in vitro applications.56−58 On running gelatin zymography on the supernatants collected from RPE cells stimulated without LPS (Figure 3a) and with 1 μg·mL−1 LPS (Figure 3b), the band corresponding to MMP-2 was present; however, the band corresponding to MMP-9 was absent. Proteolytic bands in lanes 1, 2, and 3 in Figure 3a correspond to supernatants collected from RPE cells not stimulated with LPS at 2, 6, and 24 h, respectively. The band corresponding to 2 h in lane 1 was only just visible whereas the bands in lanes 2 and 3 were more intense, indicating that the MMP-2 activity after 2 h in the nonstimulated cells was not high enough to be detected by the technique. In Figure 3b, lanes 4, 5, and 6 represent the MMP-2 activity, which was produced from supernatants of RPE cells stimulated with LPS at 2, 6, and 24 h, respectively. In this case, the band corresponding to 2 h (lane 4) was clearly visible, indicating the increase in MMP-2 activity with LPS stimulation. The increase in the band intensity along with the increase in the width of the bands with time indicated an increase in the MMP2 activity. The trend of higher MMP-2 release by RPE cells was in line with the results obtained using the PSi microsensors 9950
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where Δλcells‑LPS was the shift obtained when the sensors were present in the cell culture medium with cells only; Δλno cells+LPS corresponds to responses with cell culture medium with different LPS amounts; Δλmedia was when the sensors were present in the cell culture media only (no cells, no LPS). The PSi microsensors continue to show changes in optical response until close to 100%, where all the enzyme sensitive substrate is degraded (Figures 4 and S5). Importantly, the time over which that 100% optical response is reached is dependent on the amount of LPS used to stimulate the cells. Higher LPS levels required less time to reach the 100% optical response which is indicative of more MMPs being released by the RPE and IPE cells. Upon LPS stimulation, there was an increase in the expression level of different proteases like MMP-2 and MMP9. Since these PSi sensors were modified with peptides specific to these two enzymes only, the amounts present for MMP-2 and MMP-9 were calculated individually for the two cell types using the calibration curves generated when the sensors were incubated with different protease solutions as shown in Figure S6. Table 1 presents the amount of MMP-2 and MMP-9 detected by the PSi microsensors after 6 h of LPS stimulation. These amounts were calculated by comparing the optical responses observed from RPE and IPE cells (Figures 4 and S5, respectively) with the calibration curves presented in Figure S6. The amount of MMP-2 was calculated to be ∼2−15-fold higher than the amount of MMP-9 present in solution for RPE cells whereas, for IPE cells, the MMP-2 amounts were 1.4−3.5 times higher than the MMP-9 concentration. From the data presented in Table 1, it was estimated that, at the lowest LPS stimulation amount of 1 μg·mL−1, each RPE cell produced 25.2 ± 1.00 pg (∼1.4 μg·mL−1) of MMP-2 whereas each IPE cell produced only 4.5 ± 1.04 pg (0.6 μg·mL−1) of MMP-2 per cell. The calculated amounts of MMP-9 for the two cell types were 2.53 ± 1.45 and 1.7 ± 1.00 pg, respectively. The ability to estimate the quantity of enzyme present in solution is crucial in diseases like uveitis where proteases play an important role. For example, it has been demonstrated previously that MMP-2 levels of 6.8 ng·mL−1 in the aqueous humor of rat eyes are considered higher than normal levels leading to endotoxin-induced uveitis (EIU).59 In another study by Di Girolamo and co-workers,60 it was observed that the MMP-2 levels were higher than levels of MMP-9 in an EIU rat model for up to 12 h. However, because fluid from the aqueous humor was used in this instance, the cell type(s) secreting these proteases was not unambiguously known. The results from the PSi microsensors were also compared to a commercially available ELISA. Two different ELISA kits were used to detect the presence of the two different MMPs. The data obtained from running an ELISA on supernatants collected from LPS (1 μg·mL−1) and no LPS (0 μg·mL−1, control) RPE and IPE cells at 6 h after stimulation is presented in Figure 5. It was calculated that, on average, a single RPE and IPE cell releases 20.3 ± 1.01 and 4.14 ± 0.3 pg of MMP-2, respectively (Figure 5a), whereas much lower amounts of MMP-9 were released, 1.78 ± 0.16 pg for a RPE cell and 1.56 ± 0.06 pg by an IPE cell (Figure 5b). The results obtained between the two methods (in one method, the protease release was monitored in vitro at 6 h (using the PSi microsensors), whereas in the other method (ELISA), the supernatants collected at 6 h were added to an antibody-coated ELISA plate) were similar. These results demonstrate that the data
Figure 3. (a) Zymogram images obtained on running supernatants from RPE cells (no LPS stimulation) collected at 2, 6, and 24 h. (b) Zymogram images obtained on running supernatants from LPS stimulated RPE cells collected at 2, 6, and 24 h post stimulation. All lanes represent the band corresponding to the presence of MMP-2 (66 kDa). Active MMP-9 if detected should have been present around 84 kDa. Note: The two images were collected from two different gels.
(Figure 2a). However, the absence of MMP-9 in the zymogram demonstrates that the detection limit of PSi microsensors is lower than that obtained with zymography. Quantification of MMP-2 and MMP-9 Released by Primary Cells. Once it was confirmed that the polymer− peptide modified PSi microsensors can be used to monitor MMP-2 and MMP-9 enzymatic activity from primary cells, the next step was to quantify the amounts of MMP-2 and MMP-9 being released by RPE and IPE cells. For the detection of proteases from these two cell types, the cells were stimulated with different LPS amounts (1, 2, 5, and 10 μg·mL−1). The reflectivity measurements from the microsensors were performed in the presence of cells, as distinct from just collecting the supernatant and incubating the microsensors in the supernatant. In this way, the cells continued to release the proteases such that there was a continual supply of active MMPs. Both MMP-2 and MMP-9 measurements were performed in this manner. Each PSi microsensor was only read out once at 2, 4, 6, 8, 10, 24, and 48 h after the LPS stimulation of the cells (Figures 4 and S5). Optical response was normalized with the controls and was calculated as follows: optical reponse(%) =
Δλcells + LPS − Δλcontrols × 100 Δλpeptide + polymer
Δλcontrols = Δλcells − LPS + Δλnocells + LPS + Δλmedium
(2)
Figure 4. Optical responses obtained at different time points (2−48 h) when (a) MMP-2 and (b) MMP-9 peptide modified PSi microsensors were added to LPS stimulated RPE cells. The data is normalized to the response observed when there was no LPS added to the cells. Four different LPS concentrations were used: 1, 2, 5, and 10 μg·mL−1. 9951
DOI: 10.1021/acs.analchem.5b02529 Anal. Chem. 2015, 87, 9946−9953
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Analytical Chemistry
Table 1. Data Showing the Concentration of MMP-2 and MMP-9 Enzymes Secreted by RPE and IPE Cells after 6 h of Stimulation with Four Doses of LPS MMP-2/nM LPS, μg·mL−1 1 2 5 10
MMP-9/nM
RPEs 21.3 42.4 67.3 292
± ± ± ±
0.9 3.3 5.1 128.1
IPEs 8.6 24.7 50.7 77.3
± ± ± ±
RPEs
2.0 8.0 17.4 21.3
1.4 8.3 61.2 175
± ± ± ±
1.7 3.0 7.9 127.7
IPEs 2.5 11.8 35.6 67.4
± ± ± ±
1.6 3.2 9.0 1.7
gel electrophoresis based technique, zymography. These PSi sensors may prove to be a valuable tool as a diagnostic technique for diseases such as osteoarthritis, anterior acute uveitis, and tumors, where quantification of protease levels are critical in developing therapies for these conditions.
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ASSOCIATED CONTENT
S Supporting Information *
The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acs.analchem.5b02529. SEMs of PSi rugate filters and optical micrographs of PSi microparticles along with the schematic of the surface chemistry utilized to modify these microparticles; the optical responses observed from incubating PSi microparticles in solutions containing MMP-9 and with IPE cells (PDF)
Figure 5. Amount of (a) MMP-2 and (b) MMP-9 released by RPE and IPE cells under LPS stimulation (1 μg·mL−1) and in their native state (with no LPS stimulation) quantified from ELISA. (a) There is an upregulation of MMP-2 production by a factor of 15 and 10 for RPE and IPE cells, respectively, after LPS stimulation. (b) Upregulation of MMP-9 by a factor of 20 for RPE cells and 32 for IPE cells after stimulation with 1 μg·mL−1 LPS.
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obtained with the PSi microsensors and ELISA were concordant and are a testament of the use of PSi based microsensors as an in vitro detection technique. The slight differences in values were attributed to the differences that exist from one batch of cells to the others. The detection of low levels of enzymes (both MMP-2 and MMP-9) along with rapid and accurate protease detection in vitro allows for the development of dynamic assays using PSi microsensors for use in clinical procedures.
AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Author Contributions
The manuscript was written through contributions of all authors. B.G. designed, performed, and analyzed the experiments. K.G., P.J.R., and J.J.G. helped in designing the experiments and writing the paper. S.B.L. assisted in the analysis of the experiments shown in Figure 1 and Figure S4. K.M. cultured the primary cells used in the studies. K.M., N.D.G., and D.W. provided technical assistance for the experiments shown in Figure 3. All authors contributed toward the discussion of the results and reviewed the results presented in the manuscript.
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CONCLUSIONS In conclusion, the work presented here shows the potential of using PSi based smart surfaces as a selective and sensitive tool for detecting and quantitating proteases. The polymer−peptide modified PSi microsensors were demonstrated as sensitive optical sensors to protease enzymes in solution. Exposure to protease from nanomolar to micromolar concentrations was detected with no indication of inhomogeneous proteolysis. The enzymatic concentrations were modeled using the Michaelis− Menten kinetics, and the kinetic parameters were comparable to a solution phase assay. This suggested the rapid infiltration of the enzyme from the bulk solution into the nanoporous network along with the validation of the optical data highlighting the potential use of such a PSi based system as an interesting model to study enzyme kinetics. The selectivity of the PSi microsensors toward MMP-2 and MMP-9 demonstrated in our earlier work35 facilitated the ability of quantifying the amounts of MMP-2 and MMP-9 released by each RPE and IPE cell, which was previously unattainable through the PSi optical biosensor. Both the RPE and IPE cells produced higher amounts of MMP-2 than MMP9 when stimulated with LPS. Toward advanced optical tools for cell biology, the capability of the PSi biosensor to detect picogram levels of proteases was presented, and as a result, the sensitivity achieved for the sensor was higher than that for the
Notes
The authors declare no competing financial interest.
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ACKNOWLEDGMENTS The authors would like to thank the electron microscopy unit and biomedical imaging facility at UNSW Australia for their help and support. This research was supported by the National Health and Medical Research Council (project grant number 1024723) to J.J.G., D.W., and N.D.G. and a fellowship to K.G. (grant number 1059278).
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REFERENCES
(1) Moin, K.; Schwartz, D.; Mullins, S.; Sloane, B. In Proteases and Cancer; Bugge, T. H., Antalis, T. M., Eds.; Humana Press: New York, 2009; pp 49−57. (2) Gordon, S. Cell 2002, 111, 927−930. (3) Xia, Z.; Triffitt, J. T. Biomed. Mater. 2006, 1, R1. (4) Kumar, A.; Zhang, J.; Yu, F.-S. X. Invest. Ophthalmol. Visual Sci. 2004, 45, 3513−3522. 9952
DOI: 10.1021/acs.analchem.5b02529 Anal. Chem. 2015, 87, 9946−9953
Article
Analytical Chemistry (5) Yeh, S.; Pflugfelder, S. C.; Stern, M. E. Pathological effects of lacrimal keratoconjunctivitis on the ocular surface. In Dry eye and the ocular surfacea unified approach; Marcel Dekker: New York, 2004; Chapter 9, pp 191−203. (6) Sobrin, L.; Liu, Z.; Monroy, D. C.; Solomon, A.; Selzer, M. G.; Lokeshwar, B. L.; Pflugfelder, S. C. Invest. Ophthalmol. Visual Sci. 2000, 41, 1703−1709. (7) Li, D.-Q.; Lokeshwar, B. L.; Solomon, A.; Monroy, D.; Ji, Z.; Pflugfelder, S. C. Exp. Eye Res. 2001, 73, 449−459. (8) Eggeling, C.; Jager, S.; Winkler, D.; Kask, P. Curr. Pharm. Biotechnol. 2005, 6, 351−371. (9) Eggeling, C.; Kask, P.; Winkler, D.; Jäger, S. Biophys. J. 2005, 89, 605−618. (10) Folin, O.; Ciocalteu, V. J. Biol. Chem. 1927, 73, 627−650. (11) Knauper, V.; Lopez-Otin, C.; Smith, B.; Knight, G.; Murphy, G. J. Biol. Chem. 1996, 271, 1544−1550. (12) Kleiner, D. E.; Stetler-Stevenson, W. G. Anal. Biochem. 1994, 218, 325−329. (13) Oliver, G. W.; Leferson, J. D.; Stetler-Stevenson, W. G.; Kleiner, D. E. Anal. Biochem. 1997, 244, 161−166. (14) Yan, S. J.; Blomme, E. A. Vet. Pathol. 2003, 40, 227−236. (15) Leber, T. M.; Balkwill, F. R. Anal. Biochem. 1997, 249, 24−28. (16) Chin, V.; Collins, B. E.; Sailor, M. J.; Bhatia, S. N. Adv. Mater. 2001, 13, 1877−1880. (17) Canham, L. T. Adv. Mater. 1995, 7, 1033−1037. (18) Steinhauer, C.; Ressine, A.; Marko-Varga, G.; Laurell, T.; Borrebaeck, C. A. K.; Wingren, C. Anal. Biochem. 2005, 341, 204−213. (19) Kilian, K. A.; Lai, L. M. H.; Magenau, A.; Cartland, S.; Böcking, T.; Di Girolamo, N.; Gal, M.; Gaus, K.; Gooding, J. J. Nano Lett. 2009, 9, 2021−2025. (20) Kilian, K. A.; Böcking, T.; Gaus, K.; Gal, M.; Gooding, J. J. ACS Nano 2007, 1, 355−361. (21) Guan, B.; Magenau, A.; Kilian, K. A.; Ciampi, S.; Gaus, K.; Reece, P. J.; Gooding, J. J. Faraday Discuss. 2011, 149, 301−317. (22) Orosco, M. M.; Pacholski, C.; Sailor, M. J. Nat. Nanotechnol. 2009, 4, 255−258. (23) Orosco, M. M.; Pacholski, C.; Miskelly, G. M.; Sailor, M. J. Adv. Mater. 2006, 18, 1393−1396. (24) Bonanno, L. M.; DeLouise, L. A. Langmuir 2007, 23, 5817− 5823. (25) Bonanno, L. M.; Kwong, T. C.; DeLouise, L. A. Anal. Chem. 2010, 82, 9711−9718. (26) Bonanno, L. M. a.; DeLouise, L. A. Biosens. Bioelectron. 2007, 23, 444−448. (27) Lin, V. S. Y.; Motesharei, K.; Dancil, K. P. S.; Sailor, M. J.; Ghadiri, M. R. Science 1997, 278, 840−843. (28) Dancil, K.-P. S.; Greiner, D. P.; Sailor, M. J. J. Am. Chem. Soc. 1999, 121, 7925−7930. (29) Cunin, F.; Schmedake, T. A.; Link, J. R.; Li, Y. Y.; Koh, J.; Bhatia, S. N.; Sailor, M. J. Nat. Mater. 2002, 1, 39−41. (30) Jane, A.; Dronov, R.; Hodges, A.; Voelcker, N. H. Trends Biotechnol. 2009, 27, 230−239. (31) Gupta, B.; Zhu, Y.; Guan, B.; Reece, P. J.; Gooding, J. J. Analyst 2013, 138, 3593−3615. (32) Krismastuti, F. S. H.; Pace, S.; Melville, E.; Cowin, A.; Dargaville, T. R.; Voelcker, N. H. Aust. J. Chem. 2013, 66, 1428−1434. (33) Krismastuti, F. S. H.; Pace, S.; Voelcker, N. H. Adv. Funct. Mater. 2014, 24, 3639−3650. (34) Martin, M.; Taleb Bendiab, C.; Massif, L.; Palestino, G.; Agarwal, V.; Cuisinier, F.; Gergely, C. Physica Status Solidi (c) 2011, 8, 1888−1892. (35) Soeriyadi, A. H.; Gupta, B.; Reece, P. J.; Gooding, J. J. Polym. Chem. 2014, 5, 2333−2341. (36) Nagai, Y.; Akashi, S.; Nagafuku, M.; Ogata, M.; Iwakura, Y.; Akira, S.; Kitamura, T.; Kosugi, A.; Kimoto, M.; Miyake, K. Nat. Immunol. 2002, 3, 667−672. (37) Lan, J.; Kumar, R. K.; Di Girolamo, N.; McCluskey, P.; Wakefield, D. Br. J. Ophthalmol. 2003, 87, 208−211.
(38) Ciampi, S.; Böcking, T.; Kilian, K. A.; Harper, J. B.; Gooding, J. J. Langmuir 2008, 24, 5888−5892. (39) Hu, D. N.; Ritch, R.; McCormick, S. A.; Pelton-Henrion, K. Invest. Ophthalmol. Visual Sci. 1992, 33, 2443−2453. (40) An, E.; Lu, X.; Flippin, J.; Devaney, J. M.; Halligan, B.; Hoffman, E.; Csaky, K.; Hathout, Y. J. Proteome Res. 2006, 5, 2599−2610. (41) Di Girolamo, N.; Lloyd, A.; McCluskey, P.; Filipic, M.; Wakefield, D. Am J Pathol. 1997, 150, 653−666. (42) Bovard, B. G. Appl. Opt. 1993, 32, 5427−5442. (43) Link, J. R.; Sailor, M. J. Proc. Natl. Acad. Sci. U. S. A. 2003, 100, 10607. (44) Sailor, M. J.; Link, J. R. Chem. Commun. 2005, 1375−1383. (45) Gaspers, P. B.; Robertson, C. R.; Gast, A. P. Langmuir 1994, 10, 2699−2704. (46) Trigiante, G.; Gast, A. P.; Robertson, C. R. J. Colloid Interface Sci. 1999, 213, 81−86. (47) Kim, J.-H.; Roy, S.; Kellis, J. T.; Poulose, A. J.; Gast, A. P.; Robertson, C. R. Langmuir 2002, 18, 6312−6318. (48) Turk, B. E.; Huang, L. L.; Piro, E. T.; Cantley, L. C. Nat. Biotechnol. 2001, 19, 661−667. (49) Kridel, S. J.; Chen, E.; Kotra, L. P.; Howard, E. W.; Mobashery, S.; Smith, J. W. J. Biol. Chem. 2001, 276, 20572−20578. (50) Luo, L.; Li, D. Q.; Doshi, A.; Farley, W.; Corrales, R. M.; Pflugfelder, S. C. Invest. Ophthalmol. Visual Sci. 2004, 45, 4293−4301. (51) El-Shabrawi, Y. G.; Christen, W. G.; Foster, S. C. Curr. Eye Res. 2000, 20, 211−214. (52) Di Girolamo, N.; Verma, M. J.; McCluskey, P. J.; Lloyd, A.; Wakefield, D. Curr. Eye Res. 1996, 15, 1060−1068. (53) Hofmaier, F.; Hauck, S. M.; Amann, B.; Degroote, R. L.; Deeg, C. A. Invest. Ophthalmol. Visual Sci. 2011, 52, 2314−2320. (54) Ahir, A.; Guo, L.; Hussain, A. A.; Marshall, J. Invest. Ophthalmol. Visual Sci. 2002, 43, 458−465. (55) Eichler, W.; Friedrichs, U.; Thies, A.; Tratz, C.; Wiedemann, P. nvest. Ophthalmol. Visual Sci. 2002, 43, 2767−2773. (56) Toth, M.; Sohail, A.; Fridman, R. Methods Mol. Biol. (N. Y., NY, U. S.) 2012, 878, 121−135. (57) Sliwowska, I.; Kopczynski, Z. Wiad. Lek. (Warsaw, Poland) 2007, 60, 241−247. (58) Hawkes, S. P.; Li, H.; Taniguchi, G. T. Methods Mol. Biol. (N. Y., NY, U. S.) 2010, 622, 257−269. (59) Kalariya, N. M.; Shoeb, M.; Reddy, A. B.; Zhang, M.; van Kuijk, F. J.; Ramana, K. V. Invest. Ophthalmol. Visual Sci. 2010, 51, 5105− 5113. (60) Cuello, C.; Wakefield, D.; Di Girolamo, N. Br. J. Ophthalmol. 2002, 86, 290−295.
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