Cellulosomes are multienzyme complexes designed for digesting cellulose. AFM-based SMFS was used to study the mechanical properties and folding configuration of cellulosome-associated protein assemblies. We present a complete workflow for protein immobilization, data acquisition, and data analysis to study the interactions of individual receptor-ligand complexes involved in cellulosome assembly.
Cellulosomes are discrete multienzyme complexes used by a subset of anaerobic bacteria and fungi to digest lignocellulosic substrates. Assembly of the enzymes onto the noncatalytic scaffold protein is directed by interactions among a family of related receptor-ligand pairs comprising interacting cohesin and dockerin modules. The extremely strong binding between cohesin and dockerin modules results in dissociation constants in the low picomolar to nanomolar range, which may hamper accurate off-rate measurements with conventional bulk methods. Single-molecule force spectroscopy (SMFS) with the atomic force microscope measures the response of individual biomolecules to force, and in contrast to other single-molecule manipulation methods (i.e. optical tweezers), is optimal for studying high-affinity receptor-ligand interactions because of its ability to probe the high-force regime (>120 pN). Here we present our complete protocol for studying cellulosomal protein assemblies at the single-molecule level. Using a protein topology derived from the native cellulosome, we worked with enzyme-dockerin and carbohydrate binding module-cohesin (CBM-cohesin) fusion proteins, each with an accessible free thiol group at an engineered cysteine residue. We present our site-specific surface immobilization protocol, along with our measurement and data analysis procedure for obtaining detailed binding parameters for the high-affinity complex. We demonstrate how to quantify single subdomain unfolding forces, complex rupture forces, kinetic off-rates, and potential widths of the binding well. The successful application of these methods in characterizing the cohesin-dockerin interaction responsible for assembly of multidomain cellulolytic complexes is further described.
Cellulosomes are large multienzyme complexes displayed on the surface of anaerobic cellulolytic bacteria (e.g. C. thermocellum) that have evolved to efficiently depolymerize plant cell wall lignocellulose into soluble oligosaccharides1. A central attribute of cellulosomes is the high-affinity cohesin-dockerin interaction. In the most prominent paradigm, a highly conserved 60-75 amino acid type I dockerin module is displayed at the C-terminal end of the various bacterial enzymes. The dockerin module directs assembly of synergistic combinations of enzymes onto the noncatalytic scaffold protein ('scaffoldin'), which comprises a polyprotein of cohesin domains that are specific for the type I dockerin module. At higher levels, cellulosome architecture can become very complex, incorporating alternative cohesin and dockerin pairs (e.g. type II, type III) that anchor the structures to the cell surface and allow for the assembly of branched structures containing multiple scaffoldins2. The various cohesin-dockerin types, despite having related structures, exhibit differential binding specificities suppressing cross reactivity with unintended scaffoldins or components from other cellulosome-producing bacterial species. While bioinformatic approaches have successfully identified thousands of unique cellulosomal components at the genetic level, comparatively few protein structures are known, and the mechanisms at work in cohesin-dockerin specificity determination remains an active area of structural biology research.
Since the invention of the atomic force microscope (AFM) by Binnig et al.3, various AFM operational modes have been developed and continuously improved, including noncontact imaging, oscillation mode imaging4, and single molecule force spectroscopy (SMFS)5,6. SMFS has evolved into a widely used technique to directly probe individual proteins7-11, nucleic acids12-15, and synthetic polymers16-19. In a typical SMFS experiment to investigate receptor-ligand binding20,21, an AFM cantilever tip is modified with one of the binding partners, while a flat glass surface is modified with the complementary binding partner. The modified cantilever is brought into contact with the surface allowing the partners to bind. The base of the cantilever is then withdrawn at constant speed and the force is measured using the optical lever deflection method. The resultant force-distance data traces exhibit sawtooth-like peaks if binding was established. In cases where the binding partners are fused to multiple protein domains, each peak in the force-distance trace can be correlated to the unfolding of a single protein domain or folded subdomain, while the last peak corresponds to rupture of the protein binding interface. The specific positions of the force-resistant elements can be used as a fingerprint to identify the various protein domains of interest. This method can be used to interrogate important amino acids involved in protein folding and stabilization. Many models have been reported in the literature to treat the characteristic force extension behavior observed in SMFS experiments. The most commonly used models include the freely jointed chain (FJC) model22, the worm-like chain (WLC) model18,23-25, and the freely rotating chain (FRC) model25,26.
In our prior work11, we used single-molecule force spectroscopy to investigate the interaction of cohesin and dockerin modules. Here, we present an experimental protocol for glass surface and cantilever functionalization with enzyme-dockerin and CBM-cohesin protein constructs. We also present an AFM-based SMFS protocol including data acquisition and analysis procedures. The described protocol can easily be generalized to other molecular systems, and should prove particularly useful to researchers interested in high-affinity receptor ligand pairs.
A schematic of the pulling geometry used in this work to probe the cohesin-dockerin interaction is shown in Figure 1A. The protein immobilization protocol reported here for cantilever and cover glass functionalization is a modified version of the procedure published previously27. The proteins were expressed from plasmid vectors in E. coli using conventional methods. The proteins were designed with a solvent-accessible thiol group, which was used in combination with maleimide chemistry to tether the protein via a stable thioether linkage to the cover glass surface and cantilever. The engineered cysteine residues in both the CBM-cohesin and xylanase-dockerin fusion proteins were located towards the N-terminal side of the proteins, away from the cohesin-dockerin binding interface11. A detailed overview of the chemical bonding employed in protein immobilization is shown in Figure 1B.
1. Sample Preparation
2. Data Acquisition
In this work, a custom-built AFM28 controlled by an MFP-3D AFM controller from Asylum Research with custom written Igor Pro software was used. Cantilever deflection is measured via the optical beam deflection method29. The sample preparation and data analysis protocols provided here are applicable regardless of the exact AFM model used. However, the AFM model should be suitable for measuring in liquids and support an accessible speed range on the z-piezo of approximately 200-5,000 nm/sec.
3. Data Analysis
The flow diagram in Figure 3 illustrates the process of data analysis. Perform all data manipulations using an appropriate software package such as Igor Pro or MATLAB. First convert the raw signal from the detector into units of force, and correct for offset and drift. Subsequently, use models of biopolymer elasticity to locate energy barriers in the unfolding pathways, and identify protein subdomains. Finally, kinetic and energetic parameters of the receptor-ligand interaction are obtained.
We used the described procedure to investigate a type I cohesin-dockerin pair from C. thermocellum. Upon successful binding of the cohesin-dockerin pair, the recorded force distance traces showed characteristic peak patterns. A typical trace is shown in Figure 4a. Every peak in the trace represents the unfolding of one protein subdomain with the last peak corresponding to the dissociation of the receptor-ligand complex.
For the CBM-cohesin-dockerin-xylanase complex investigated in this work, the initial rise in force corresponds to stretching of the PEG linker molecules. The subsequent series of up to three descending force dips reflects the unfolding of the xylanase domain. The final peak represents the rupture of the cohesin-dockerin binding interface.
All recorded force-distance traces were transformed to force-contour length space. The resulting barrier position histogram is shown in Figure 4B. The data show a contour length increment of approximately 89 nm. The xylanase domain consists of 378 amino acids, 260 of which are located C-terminally from the engineered cysteine residue. From the crystal structure, the folded length of the domain is assumed to be 6 nm. Further assuming a length per stretched amino acid of 0.365 nm35, the measured 89 nm increment can be unambiguously assigned to the unfolding of the xylanase domain. This is consistent with previously published results11.
To probe the energy landscape of the cohesin-dockerin interaction, we analyzed a total of 186 data traces obtained with four different pulling speeds (0.2, 0.7, 2.0, and 5.0 µm/sec). The resulting force spectrum is shown in Figure 4C. Fitting Equation (5) to the data yields values for koff and Dx of 3.13 x10-5/sec and 0.70 nm, respectively. These values are in good agreement with previously published results11.
Figure 1. Schematic of biomolecule immobilization. (A) Xylanase-dockerin fusion proteins are attached to the glass slide via PEG linkers. The cantilever is similarly modified with a cohesin protein fused to a cellulose binding module (CBM). (B) Depiction of chemical bonding employed in cover glass and cantilever functionalization.
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Figure 2. Process flow diagram showing sample preparation steps followed by data acquisition and analysis.Click here to view larger image.
Figure 3. Data analysis workflow diagram showing the processing steps involved in converting the raw detector signals into force-extension traces. These traces are further analyzed to obtain information about receptor-ligand binding. The final results provide energetic and kinetic parameters about specific domains.
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Figure 4. Single molecule force spectroscopy data on cohesin-dockerin. (A) Typical unfolding trace showing PEG linker stretching, xylanase unfolding, and rupture of the cohesin-dockerin binding interface. (B) Contour length histogram assembled from 314 force distance traces exhibiting energy barrier positions along the contour length. (C) Dynamic force spectrum obtained from 186 force-extension traces. Large blue circles represent the most probable rupture force at a given loading rate. The solid line represents a least squares fit to Equation 5. Rupture event populations are shown in the background. Error bars represent standard deviation obtained from Gaussian fits.
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To obtain meaningful data from single molecule force spectroscopy experiments, it is crucial to achieve well-defined and reproducible pulling geometries. The protocol used here results in site-specific immobilization of protein complexes in a defined pulling geometry.
The cantilevers used in this study were chosen due to their force sensitivity and high resonance frequency in water. Moreover, the small tip curvature of approximately 10 nm is advantageous for single molecule experiments due to reduced likelihood of multiple interactions. However, the small footprint (38×16 µm2) of the cantilever arm complicates the adjustment of the laser beam when the optical deflection method29 is used. The diameter of the focused laser beam in the setup used for this study is comparable to the width of the cantilever. As a result, obtaining a steady sum signal can be difficult. The laser drift on the cantilever can be partially compensated for using noise analysis across the data curves to correct the inverse optical lever sensitivity, as we have described. A new atomic force microscope with a shortened optical path and smaller laser spot size is currently under development in our group to improve data quality.
To obtain reliable information about rupture events, analysis of many traces is necessary. Since single molecule force spectroscopy measurements are subject to various fluctuations, averaging in force-extension space is not constructive. Barrier position histograms, however, once aligned in contour length space can be averaged since they are independent of fluctuations. As a result, even tiny features in the unfolding pathway are resolved. Conventional superposition of force extension traces does not achieve this kind of resolution.
In a force regime above 500 pN, a corrected WLC model accounting for electron cloud elasticity (QM-WLC) describes force-extension behavior better than the classical WLC model18. The freely rotating chain26 model (FRC) can also be used in a high force regime. With rupture forces up to 125 pN, the cohesin-dockerin interface shows one of the strongest receptor-ligand interactions reported in the literature. The WLC model was used in this work and in practice there was little difference between WLC, QM-WLC, and FRC models for analysis of cohesin-dockerin unfolding traces.
The conventional Bell-Evans20,34 model was used to analyze the force-loading rate dependency of the cohesin-dockerin binding interface. Recent works36,37 have extended the theoretical framework for the interpretation of single molecule experiments. These models treat nonlinear trends in the force spectra. Furthermore, they produce the free energy barrier height DG of the dissociation event. To observe distinct nonlinear trends in the force spectra, loading rates need to be varied over many orders of magnitude. Realizing extremely low loading rates is theoretically achievable using extremely slow z-piezo pulling speeds, however in practice this poses a challenge due to drift in the tip-substrate distance. Extremely high loading rates can also be difficult to obtain since increasing noise might obscure certain features in the recorded force-distance traces. Choice of the theoretical model must be balanced with these practical aspects of data acquisition while considering the specific proteins under investigation. In many cases the linear Bell-Evans model is entirely sufficient.
In conclusion, a complete experimental protocol for the study of receptor-ligand interactions using AFM-based single-molecule force spectroscopy has been presented. The positioning accuracy and force sensitivity of the atomic force microscope in conjunction with versatile biomolecule immobilization strategies provide an excellent toolbox for the investigation of receptor-ligand systems for structural biology studies.
The authors have nothing to disclose.
The authors acknowledge funding from a European Research Council advanced grant to Hermann Gaub. Michael A. Nash gratefully acknowledges funding from Society in Science – The Branco Weiss Fellowship program. The authors thank Edward A. Bayer, Yoav Barak, and Daniel B. Fried at the Weizmann Institute of Science for generously providing the proteins used in this study. The authors thank Hermann E. Gaub, Elias M. Puchner, and Stefan W. Stahl for helpful discussions.
3-Aminopropyl dimethyl ethoxysilane | ABCR GmbH | AB110423 | |
5 kDa NHS-PEG-maleimide | Rapp Polymer | 13 5000-65-35 | |
TCEP Disulfide reducing gel | Thermo Scientific, Pierce | 77712 | www.thermoscientific.com/pierce |
Tris(hydroxymethyl)aminomethane | |||
BioLever mini silicon nitride cantilevers | Olympus | BL-AC40TS-C2 | Soft batches |
XYZ Piezoelectric actuators | Physik Instrumente GmbH | ||
Infrared “broad spectrum” IR laser | Superlum | ||
MFP-3D AFM Controller | Asylum Research | ||
Igor Pro 6.31 | Wavemetrics | Data acquisition and analysis | |
Sodium chloride | |||
Calcium chloride | |||
pH Meter | |||
Sodium borate | |||
Tweezers | |||
Cover glasses | Thermo Scientific, Menzel-Gläser | 24 mm diameter, 0.5 mm thickness | |
PTFE sample holder | custom made | ||
Sonicator bath | |||
Ethanol | analytical purity | ||
Sulfuric acid (concentrated) | analytical purity | ||
Hydrogen peroxide (30%) | analytical purity | ||
Orbital shaker | |||
Toluene | analytical purity | ||
Filter paper | |||
Glass slides | |||
Microtubes | |||
Micropipettes | |||
Centrifuge | suitable for microtubes | ||
Rotator | |||
Petri dishes | |||
Beakers | |||
Optical microscope |