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Enhancing Coupled Enzymatic Activity by Colocalization on Nanoparticle Surfaces: Kinetic Evidence for Directed Channeling of Intermediates James Nicholas Vranish,†,‡,# Mario G. Ancona,§ Eunkeu Oh,∥,⊥ Kimihiro Susumu,∥,⊥ Guillermo Lasarte Aragonés,‡,¶ Joyce C. Breger,‡ Scott A. Walper,‡ and Igor L. Medintz*,‡ †
National Research Council, Washington, DC 20001, United States Center for Bio/Molecular Science and Engineering, Code 6900, §Electronic Science and Technology Division, Code 6800, and ∥ Optical Sciences Division, Code 5611, U.S. Naval Research Laboratory, Washington, DC 20375, United States ⊥ KeyW Corporation, Hanover, Maryland 21076, United States # Department of Chemistry and Physics, Ave Maria University, Ave Maria, Florida 34142, United States ¶ College of Science, George Mason University, Fairfax, Virginia 22030, United States ‡
S Supporting Information *
ABSTRACT: Multistep enzymatic cascades are becoming more prevalent in industrial settings as engineers strive to synthesize complex products and pharmaceuticals in economical, environmentally friendly ways. Previous work has shown that immobilizing enzymes on nanoparticles can enhance their activity significantly due to localized interfacial effects, and this enhancement remains in place even when that enzyme’s activity is coupled to another enzyme that is still freely diffusing. Here, we investigate the effects of displaying two enzymes with coupled catalytic activity directly on the same nanoparticle surface. For this, the well-characterized enzymes pyruvate kinase (PykA) and lactate dehydrogenase (LDH) were utilized as a model system; they jointly convert phosphoenolpyruvate to lactate in two sequential steps as part of downstream glycolysis. The enzymes were expressed with terminal polyhistidine tags to facilitate their conjugation to semiconductor quantum dots (QDs) which were used here as prototypical nanoparticles. Characterization of enzyme coassembly to two different sized QDs showed a propensity to cross-link into nanoclusters consisting of primarily dimers and some trimers. Individual and joint enzyme activity in this format was extensively investigated in direct comparison to control samples lacking the QD scaffolds. We found that QD association enhances LDH activity by >50-fold and its total turnover by at least 41-fold, and that this high activation appears to be largely due to stabilization of its quarternary structure. When both enzymes are simultaneously bound to the QD surfaces, their colocalization leads to >100-fold improvements in the overall rates of coupled activity. Experimental results in conjunction with detailed kinetic simulations provide evidence that this significant improvement in coupled activity is partially attributable to a combination of enhanced enzymatic activity and stabilization of LDH. More importantly, experiments aimed at disrupting channeled processes and further kinetic modeling suggest that the bulk of the performance enhancement arises from intermediary “channeling” between the QDcolocalized enzymes. A full understanding of the underlying processes that give rise to such enhancements from coupled enzymatic activity on nanoparticle scaffolds can provide design criteria for improved biocatalytic applications. KEYWORDS: enzyme, nanotechnology, nanoparticle, quantum dot, catalysis, kcat, enhancement, synthetic biology, biocatalysis, substrate channeling
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nzymes are well-known for their extraordinary catalytic properties. They can enhance the rate of reactions that would otherwise require extreme conditions to © XXXX American Chemical Society
Received: March 28, 2018 Accepted: July 12, 2018
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DOI: 10.1021/acsnano.8b02334 ACS Nano XXXX, XXX, XXX−XXX
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Figure 1. PykA−LDH enzyme cascade. (A) Reaction scheme for coupled PykA and LDH activity. Individual and coupled enzyme activity is monitored by NADH consumption and loss of its absorption at 340 nm. (B) Schematic of the coupled PykA−LDH enzyme system colocalized on a QD surface. The propensity of the enzymes to form cross-linked QD dimers and, to a lesser extent, trimers via the enzymes tetrameric polyhistidine tags is also schematically indicated. Note, not to scale.
synthesis is being pursued concurrently as an alternative approach to achieving the same goals.4−8,10,11 The latter modality is attractive for directly bypassing reliance on cells and reducing the system to its core-minimal components, but cell-free or in vitro approaches also have their own challenges. Among the issues are the need to maintain the enzymes in a viable state, to regenerate essential cofactors, and to overcome diffusion limitations once the tight confines of the cellular microenvironment have been removed.12 Currently, cell-free implementation often involves the immobilization and colocalization of the necessary enzymes on some type of surface or carrier,13−16 as this is often found to reliably stabilize the enzymes.15,17,18 However, in many cases, such immobilization also degrades the enzyme’s activity, and this represents a primary technological roadblock toward achieving highly efficient cell-free biosynthesis.19−21 Why an enzyme’s activity decreases following immobilization on a surface is still not fully understood and has been variously ascribed to a complex interplay of contributing factors. The latter include the chemistry used to immobilize the enzymes which can modify necessary functional groups on key reactive residues, nonproductive orientation of the enzyme especially when using heterogeneous attachment chemistries,
accomplish while simultaneously offering unmatched control over reaction site selectivity and stereospecificity.1 The power of single enzymes is hugely magnified in biology by deploying them in targeted cascades in which multiple enzymes act in concert to biocatalyze the synthesis of incredibly complex products, many of which still cannot be replicated in vitro. Due to these benefits, enzymes are frequently utilized in industry and especially in pharmaceutical synthesis.2,3 Given their biological origin, assembling and exploiting enzymes within host organisms for chemical production by metabolic engineering with the goal of deriving key industrial and drug precursors has garnered strong research interest in the nascent field of synthetic biology.4−7 Although potentially revolutionary, the use of such living cellular foundries faces many challenges. These include the complexity of recombinantly engineering non-native enzymatic pathways or cascades in a given cellular expression host; the high energy demands and delicate environmental balance that must be maintained to keep cells alive; substrate and product transport into and out of cells, respectively, across a membrane barrier; the presence of many competing and parasitic enzyme pathways; and intermediary, product, or even direct enzyme toxicity.7−9 Given these issues, development of cell-free enzymatic B
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RESULTS Enzymes, Assays, and Quantum Dots. A detailed description of the materials and enzyme preparation along with the enzymatic assay formats is provided in the Methods and Supporting Information (SI). The genes for bacterial Dlactate dehydrogenase (LDH, EC 1.1.1.28) and pyruvate kinase (PykA, EC 2.7.1.40) were cloned directly from Escherichia coli strain BL21(DE3) and inserted into the pET28b vector to add an N-terminal (His)6 tag to each. Both proteins were expressed in E. coli strain NiCo21(DE3) and isolated to >90% purity using immobilized metal affinity chromatography (IMAC) (see SI Figure S3 for representative electropherograms). The enzyme molecular weights (MWs) as shown by a Bio-Rad protein chip assay (39.1 and 56.1 kDa for LDH and PykA, respectively) correspond well with the predicted masses of 38.7 and 53.5 kDa for each of their monomeric units. X-ray crystal structures and gel-filtration chromatography studies indicate that the enzymes are both homotetramers,37,38 with approximate MWs of ∼160 (LDH) and ∼220−240 kDa (PykA). As each monomer has a terminal (His)6 tag, each homotetramer should therefore have 4 × (His)6, which can allow them to bind multiple QDs, as was previously shown for β-galactosidase.32 Due to its functional ramifications, this aspect is addressed in depth further below. Enzyme units and activity referred to in the text are typically for the number of monomers per QD unless otherwise specified as tetramers per QD in certain circumstances. Both enzymes provide catalytic activity that is an integral part of a cell’s ability to derive energy from glucose via glycolysis. As shown in Figure 1A, PykA utilizes phosphoenolpyruvate (PEP) as a phosphate donor to phosphorylate adenosine diphosphate (ADP) substrate and regenerates adenosine triphosphate (ATP) while yielding pyruvate. LDH, in turn, converts pyruvate to lactate using reduced nicotinamide adenine dinucleotide (NADH) as the hydride donor/cofactor leaving oxidized NAD+.39 As described in the Methods, PykA activity is assayed by monitoring pyruvate product generation using the coupled activity of LDH from Lactobacillus leichmanii (LL-LDH) and the loss of NADH absorption at 340 nm under conditions where NADH and LLLDH are not rate-limiting. LL-LDH does not display a (His)6 and thus will not coordinate the QDs in these assays. Moreover, we did not observe any change in LL-LDH activity in the presence/absence of QD. LDH and coupled PykA → LDH activity were also directly assayed by monitoring loss of NADH absorption. Assays were performed in 384-well plates in parallel using a microtiter plate reader, as described in the Methods, in formats where the enzymes were either selfassembled onto the QDs (Figure 1B) or with the equivalent amount of freely diffusing enzyme (no QD controls) in identical volumes. Although we have utilized gold NPs as scaffolds for similar types of studies,40 we most often utilize nanocrystalline CdSe/ ZnS core/shell QDs due to the specific benefits they provide for these types of enzymatic studies.30−32,41−44 In particular, the QDs can be prepared with tight control over size (polydispersity ≤10%) and can be colloidally dispersed in aqueous media using a wide variety of stabilizing ligands that provide different charges, polarities, and steric bulk to the QD surfaces as desired.45−48 Use of such differing ligands has already been shown to dramatically influence protease activity on QD−substrate constructs.48 QDs are well-known for their
and the nature and size of the surface which can favor laminar flow and limit the diffusional mixing needed for optimal activity. A growing suite of available bioorthogonal chemistries is now able to partially address attachment issues by providing chemical modifications that are both site-specific and provide for oriented enzyme−surface binding.22−24 In terms of carrier choice, a variety of materials are currently being explored, ranging from particles to cellular surfaces and other biologicals such as engineered virions.25−28 Within this library of potential scaffolds, nanoparticles (NPs), in particular, have shown perhaps the most promise, as displaying enzymes around them very often results not in a depression of enzymatic activity but rather, counterintuitively, an enhancement.19,25,29 Kinetic enhancement following NP display has been verified for a plethora of diverse enzyme types and sizes including alkaline phosphatase, β-galactosidase, horseradish peroxidase, recombinant esterases, etc. (see refs 10, 19, 25, and 29−32 and therein). Here, too, the exact mechanism(s) that gives rise to enhancement is still not fully understood but has been suggested to be associated with controlling the orientation of enzyme display on the NP, stabilization of structure and activity at low concentrations, alleviation of rate-limiting steps such as enzyme−product dissociation, NP interfacial or boundary effects, and/or some type of localized substrate accumulation.11,19,25,29−32 More pertinently, we have recently shown that enhancement can be maintained in coupled catalytic systems (i.e., where the product of one enzyme is the substrate of the other) where one enzyme was attached to a NP and the other remained freely diffusing.10 When horseradish peroxidase was attached to semiconductor quantum dots (QDs), its activity was both stabilized and enhanced with the observed enhancement maintained even when it was coupled to the activity of freely diffusing glucose oxidase. Going beyond this with NP coassembly of enzymes that engage in coupled activity brings with it another interesting question beyond just that of individual rate enhancements, that is, the possibility for substrate or intermediary channeling, especially given the nanoscale confines of the localized environment.19,33−35 When effectively implemented, substrate channeling can augment the activity of coupled enzymatic steps by orders of magnitude.36 Here, we describe a critical step toward assembling a biosynthetic enzyme cascade on a NP by evaluating the activity of two coupled enzymes colocalized on the same NP surface. For these purposes, the coupled glycolytic enzymes pyruvate kinase (PykA) and lactate dehydrogenase (LDH) which participate in some of the downstream steps of glycolysis were conjugated to a QD surface (see Figure 1). The impact of QD conjugation on their individual activities was first measured, and the origin of a >50-fold enhancement in LDH kinetic activity was identified. The kinetics of their combined activities when bound to a QD surface was then characterized, with greater than 100-fold enhancement in specific activity observed relative to the same concentrations of freely diffusing control samples. Finally, a series of focused studies in combination with detailed kinetic modeling were undertaken to understand the mechanism(s) that gives rise to the enhanced activity and to assess whether “channeling” of pyruvate between the two enzymes colocalized on a QD contributes to the observed rate enhancements. C
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Figure 2. Assembly of PykA and LDH to QDs. (A) TEM micrographs of 525 nm emitting QDs with an average diameter of 4.3 ± 0.6 nm. (B) TEM micrographs of 605 nm QDs with a length × width of 10.1 ± 1.0 nm × 4.5 ± 0.4 nm, respectively, which gives rise to an aspect ratio of 2.3 ± 0.3. Values are determined from measurements of ≥100 QDs. Inset shows a high-resolution TEM of each QD. (C) Gel image of the indicated ratios of either PykA or LDH assembled to the 525 QDs and then separated in a 1.25% agarose gel in 1× TBE buffer. The white arrows in both images indicate the newly formed band that form with enzyme binding to QDs. (D) Gel images of the indicated ratios of PykA and LDH tetramer assembled to the 525 QDs and then separated in a 1% agarose gel in 1× TBE buffer. Images captured after electrophoresis for 5 or 10 min. The ratios of PykA and LDH assembled on the QDs correspond to those in Figure 4B,C. Results from dynamic light scattering analysis plotting hydrodynamic diameter (HD) versus the indicated (E) PykA or (F) LDH ratios assembled to each QD sample. Note, enzyme ratios are the number of tetramers per QD here. (G) TEM micrographs of 605 nm emitting QDs assembled at an 8:4:1 mixture of LDH/PykA/QD, with arrows highlighting some of the enzyme cross-linked QD clusters at (i) 100 nm, (ii) 20 nm, and (iii) 10 nm scale resolution. QDs with smaller diameter are perpendicular to the plane of view. Note the difference in appearance and clumping as compared to that in the QD-only samples in panels A and B.
images of the CL4-functionalized 525 and 605 QD samples utilized in this study. The 525 QDs have a roughly spherical shape with an average diameter of 4.3 ± 0.6 nm, whereas the 605 QDs have a length × width of 10.1 ± 1.0 nm × 4.5 ± 0.4 nm for an aspect ratio of 2.3 ± 0.3. As described in the SI, the maximum enzyme loading on these QDs is roughly estimated to be 1 or 2 PykA tetramers and 9 or 12 LDH tetramers per 525 QD and 605 QD, respectively.52 The ability of both LDH and PykA to self-assemble to the surface of the QDs by metal affinity coordination was assessed with agarose gel shift assays.53 The representative images in Figure 2C show the shifts resulting from increasing ratios of either enzyme when assembled on the 525 QDs and separated on a 1.25% w/v agarose gel. For both enzymes, the appearance of a newly formed discrete QD conjugate band displaying significantly slower migration is apparent, and this band becomes more pronounced as the ratio of enzyme assembled increases. The conjugate band first becomes visible at a ratio of ∼0.5
size-tunable optical properties, and these have also been exploited for FRET-based sensing of enzymatic activity.48−50 The property of most relevance to the current experimental format is the ability to ratiometrically conjugate QDs by selfassembly with discrete numbers of biomolecules, such as enzymes, while also having control over orientation (in the case of monomeric proteins) in a facile manner using metal affinity coordination. This is based on the strong interaction between the enzyme’s terminal (His)6 motif and the QD’s ZnS surface. This bioconjugation is nearly spontaneous, with high affinity (Kd ∼ 1 × 10−9 M), arises from multiple synergistic interactions, and requires only mixing of the two constituents in the desired molar ratios.51 525 nm emitting (green) and 605 nm emitting (red) QDs are used in the current study and are dispersed in buffer using compact ligand CL4 (see SI Figure S1 for the chemical structure).45 Characterization of PykA and LDH Assembly to QDs. Figure 2A,B shows transmission electron microscopy (TEM) D
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Figure 3. Kinetic characterization of PykA−QD and LDH−QD conjugates. (A) Fold rate enhancement of PykA monomers assembled on the QDs at the indicated ratios plotted as a function of the PykA to QD ratio. Enhancement is as compared to the equivalent amount of free enzyme control. Reactions contained 1.56 nM PykA, 5 mM MgCl2, 1.19 mM ADP, 200 μM NADH, 7.2 mM PEP, and saturating concentrations of commercial LDH. Ratio of enzyme to QD was altered by varying the QD concentration. The specific activity (S.A., measured as units of μM NADH consumed·s−1·μM enzyme−1) of PykA was measured as a function of (B) [ADP] or (C) [PEP]. These reactions contained 1.25 nM PykA, 0.31 nM QD (4 PykA/QD), 5 mM MgCl2, 200 μM NADH, and saturating LL-LDH. Additionally, the concentration of PEP was fixed at 7.2 mM in panel B, and ADP concentration was fixed at 1.19 mM in panel C. Solid black lines represent the fits of the data to the Michaelis−Menten equation, with parameters shown in Table 1. (D) Fold rate enhancement of LDH monomers assembled on the QDs at the indicated ratios plotted as a function of the LDH to QD ratio. Enhancement is as compared to the equivalent amount of free enzyme control. Reactions contained 3.13 nM LDH, 5 mM MgCl2, 200 μM NADH, and 9 mM pyruvate. The specific activity of LDH (ratio of 1 LDH tetramer/QD) was measured as a function of (E) [NADH] or (F) [pyruvate] (measured as units of μM NADH consumed·s−1·μM enzyme−1). Lines in panels A,D−F are for visualization purposes and do not reflect a fit to the data.
increased up to 10. Considering the size of the PykA (∼13 × 9 × 11 nm from PDB entry 3T05) and LDH (9 × 7 × 10 nm from PDB entry 3WX0) homotetramers,37,38 the data confirm that the proteins are indeed assembling around the QDs. The size change in the QDs upon multiple enzyme binding is ∼20 nm, which is consistent with the addition of PykA or LDH tetramers around the NP surface. The loading of 525 QD appears to saturate with PykA between 5 and 10 equiv. of PykA tetramer. The LDH−QD combinations do not appear to be fully saturated at these ratios, and these results are consistent with our predicted loading ratios. Furthermore, when the QDs were incubated with 5 equiv. of both LDH and PykA tetramers simultaneously, the HD values were 38 ± 2 and 38 ± 3 nm for the 605 and 525 QD, respectively, which suggests that the assembled protein’s size now dominates the HD of the conjugates. This is also larger than the HD of either QD when incubated with 5 equiv. of only one of the two enzymes, again serving to confirm assembly of both to the QD surface. In addition to the initial QD-conjugated species that we observed with DLS, we also noted the formation of some larger sized entities when the QDs were incubated with enzyme, possibly indicating the presence of cross-linked QD clusters (SI Figure S4); this is also presented schematically on the right in Figure 1B. The actual amount of the aggregated materials present was hard to directly assess from the DLS data.54 To address this further, we carried out TEM imaging of the mixed enzyme samples. Representative images of the 8:4:1 mixture of
tetramers/QD for PykA and LDH. Moreover, the intensity of the native QD band decreases with the formation of this QD conjugate band which, in turn, becomes more intense with increasing ratio, as well. The native or free QD band appears to be almost gone at ratios of 10 enzyme tetramers per QD, whereas that of the QD−enzyme bioconjugate is strongest here. Similar results were obtained for the 605 QDs (data not shown). It is important to note that the predicted net charges of PykA and LDH in the TBE buffer pH 8.3 used with the gels are −4.7 and −12.4, respectively. As such, they are also expected to migrate strongly toward the anode along with the QDs. Taking this into consideration along with each protein’s homotetramic nature makes it hard to evaluate the gel data beyond confirming that the proteins are indeed self-assembling to the QDs in a manner that tracks with increased enzyme ratio. Additionally, in Figure 2D, experiments were carried out to assess the ability of both LDH and PykA to simultaneously bind to the QDs. The QD’s migration in the sample containing both enzymes is clearly slowed further than that in either sample containing just a single enzyme, indicating that both enzymes are capable of binding to the QDs simultaneously. For further characterization of the QD−enzyme conjugates, dynamic light scattering (DLS) analysis was carried out as the ratio of PykA or LDH tetramer assembled per QD was incrementally increased. As shown in Figures 2E,F, the hydrodynamic diameter (HD) of each QD sample increased around 4-fold as the ratio of enzyme tetramer to QD was E
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ACS Nano Table 1. Selected Kinetic Parameters Estimated from Experimental Dataa PykA kinetic constants sample
enzyme monomer/QD ratio
PykA PykA on 525 QD PykA on 605 QD
N/A 4 4
sample
enzyme monomer/QD Ratio
LDH LDH on 525 QD LDH on 605 QD
N/A 4 4
kcat (s−1)b
277 ± 9 214 ± 6 132 ± 6 LDH kinetic parameters max. specific activity (s−1)c
∼11 ∼250 ∼110 LDH kinetic and thermodynamic properties
KMPEP (μM)
KMADP (μM)
690 ± 70 920 ± 80 840 ± 70
130 ± 10 140 ± 10 110 ± 20
K0.5NADH (μM)d e
nd ∼800 ∼2000
K0.5pyruvate (μM)d ∼4000 ∼1800 ∼1800
sample
LDH monomer/QD ratio
kcat1 (s−1)
kcat2 (s−1)
KDapp (nM)
nH
no QD 525 QD 605 QD
N/A 4 4
45 ± 3 80 ± 20 180 ± 15
66 ± 10 0 ± 19 4±1
11.7 ± 0.8 0.7 ± 0.1 0.90 ± 0.06
4±1 3±1 6±2
Note: enzyme-only samples had the same concentration as those with QDs. bValues were obtained by fitting the ADP dependence data. cValues are the maximum specific activity observed from the pyruvate dependence data. dEstimates were obtained by interpolation of data to find the concentration that gave approximately half-maximal specific activity. eThis value was not determined due to poor signal-to-noise ratio at these low rate values; nH is the Hill coefficient. a
the QD, with the change largest for the 605 QD. The on-QD KM value for PEP is slightly reduced by 20−30%, whereas the on-QD KM value for ADP remains essentially unaffected with ≤15% variation (see Figures 3B,C and Table 1). The effect on activity of assembling LDH on QDs was investigated next. In doing so, it should be noted that the activity (structure) of LDH was enhanced by preincubation in ethylenediaminetetraacetic acid (EDTA) prior to dilution into the assay (perhaps due to protein chelation of Ni2+ that remained bound after purification; similar phenomena have been noted for stabilizing the structure of other enzymes),55 and this step was included in all studies (SI Figure S6). At low ratios of LDH to QD (5 PykA monomers (>1.25 tetramers) per QD, the inhibition appears to be alleviated and the kinetics are essentially indistinguishable from the activity of the free enzyme. We note that this trend is opposite to what we have observed previously with other enzymes.10,30−32,43 Analyses of the PykA activity both on and off the QDs were also undertaken to estimate the effective Michaelis−Menten parameters. Figure 3B,C displays the response of PykA to varying concentrations of either ADP or PEP, whereas Table 1 presents the estimated catalytic parameters. Fits to the data show a ca. 20−50% reduction in the kcat as a result of being on F
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ACS Nano catalyze each reaction as expected and formed the correct product in a manner identical to the native enzymes; and (ii) QDs alone were not capable of converting any of the substrate to product (see SI Figure S13). Mechanism of LDH Enhancement. Given the significant enhancement in LDH turnover that results from it being attached to a QD, it was important to understand the mechanistic origin(s) behind this increase before proceeding. A first clue comes from direct assays of LDH activity which found that, as the enzyme concentration increased, the specific activity did also (SI Figure S7 Although LDH is nominally a tetramer, it is known to undergo subunit dissociation at low LDH concentrations,56 and our experimental results could thus be explained by the reasonable hypothesis that the activity falls off as a result of enzyme dissociation. Furthermore, this is consistent with previous studies showing that the LDH monomer/tetramer equilibrium is reversible, but that the monomeric state can transition to an inactive species57 Indeed, the crystal structure of LDH is consistent with this conjecture as the enzyme has two of its N termini (where the His6 tags are located) paired in close proximity (∼3.5 nm) on opposite sides of the tetramer; this presumably favors the attachment of 2 (and possibly 4) His6 tags from a single LDH tetramer to a single QD nanocrystal, which would serve to effectively hold the tetrameric structure together (SI Figure S8A).37 This is in contrast to the structure of PykA, where the N-termini are >5 nm apart and buried in a cleft of the enzyme tetramer (SI Figure S8B,C).38 To test this hypothesis, the specific activity of LDH both freely diffusing and when assembled to the two QD samples was measured as a function of LDH concentration (Figure 4A). The resulting data were fit to eq 1 (Methods). Free LDH undergoes an apparent highly cooperative dissociation with a KDapp of ∼12 nM (Table 1). In contrast, the enzymes bound to either the 605 or 525 QDs dissociate in a cooperative manner, with a KDapp of
