Robust membrane protein tweezers reveal the folding speed limit of helical membrane proteins

  1. Seoyoon Kim
  2. Daehyo Lee
  3. WC Bhashini Wijesinghe
  4. Duyoung Min

  • Curated by eLife

    eLife logo

    eLife assessment

    This work presents an important methodological advance for single-molecule force spectroscopy of membrane proteins providing a new robust design of the linkage between a target single molecule and solid support. The data provide compelling evidence of the improved mechanical stability of the pulling system. Allowing more statistically reliable force measurements of biomolecules, this method may be broadly applicable in the field of single-molecule biophysics.

Reviewed by

Read the full article See related articles

Listed in

Log in to save this article

Abstract

Single-molecule tweezers, such as magnetic tweezers, are powerful tools for probing nm-scale structural changes in single membrane proteins under force. However, the weak molecular tethers used for the membrane protein studies have limited the observation of long-time, repetitive molecular transitions due to force-induced bond breakage. The prolonged observation of numerous transitions is critical in reliable characterizations of structural states, kinetics, and energy barrier properties. Here, we present a robust single-molecule tweezer method that uses dibenzocyclooctyne cycloaddition and traptavidin binding, enabling the estimation of the folding ‘speed limit’ of helical membrane proteins. This method is >100 times more stable than a conventional linkage system regarding the lifetime, allowing for the survival for ~12 hr at 50 pN and ~1000 pulling cycle experiments. By using this method, we were able to observe numerous structural transitions of a designer single-chained transmembrane homodimer for 9 hr at 12 pN and reveal its folding pathway including the hidden dynamics of helix-coil transitions. We characterized the energy barrier heights and folding times for the transitions using a model-independent deconvolution method and the hidden Markov modeling analysis, respectively. The Kramers rate framework yields a considerably low-speed limit of 21 ms for a helical hairpin formation in lipid bilayers, compared to μs scale for soluble protein folding. This large discrepancy is likely due to the highly viscous nature of lipid membranes, retarding the helix-helix interactions. Our results offer a more valid guideline for relating the kinetics and free energies of membrane protein folding.

Article activity feed

  1. Version published to 10.7554/elife.85882 on eLife
  2. eLife

    eLife assessment

    This work presents an important methodological advance for single-molecule force spectroscopy of membrane proteins providing a new robust design of the linkage between a target single molecule and solid support. The data provide compelling evidence of the improved mechanical stability of the pulling system. Allowing more statistically reliable force measurements of biomolecules, this method may be broadly applicable in the field of single-molecule biophysics.

  3. eLife

    Reviewer #1 (Public Review):

    This manuscript features a key technical advance in single-molecular force spectroscopy. The critical advance is to employ a click chemistry (DBCO-cycloaddition) for making a stable covalent connection between a target biomacromolecule and solid support in place of conventional antigen-antibody binding. This tweak dramatically improves the mechanical stability of the pulling system such that the pulling/relaxation can be repeated up to a thousand times (the previous limit was a few hundred cycles at best). This improvement is broadly applicable to various molecular interactions and other types of single-molecule force spectroscopy allowing for more statistically reliable force measurements. Another strength of this method is that all conjugation steps are chemically orthogonal (except for Spy-catcher conjugation to the termini of a target molecule) such that the probability of side reactions could be reduced.

    The reliability of kinetic and thermodynamic parameters obtained from single-molecule force spectroscopy depends on statistics, that is, the number of pulling measurements and their distribution. By extending the number of measurements, this robust method enables fundamental/critical statistical assessment of those parameters. That is, it is an important and interesting lesson from this study that ~200 repeats can yield statistically reasonable parameters.

    The authors carried out carefully designed optimization steps and inform readers of the critical aspects of each. The merit, quality, and rigor as a method-oriented manuscript are impressive. Overall, this is an excellent study.

  4. eLife

    Reviewer #2 (Public Review):

    In this study, the authors have developed methods that allow for repeatedly unfolding and refolding a membrane protein using a magnetic tweezers setup. The goal is to extend the lifespan of the single-molecule construct and gather more data from the same tether under force. This is achieved through the use of a metal-free DBCO-azide click reaction that covalently attaches a DNA handle to a superparamagnetic bead, a traptavdin-dual biotin linkage that provides a strong connection between another DNA handle and the coverslip surface, and SpyTag-SpyCatcher association for covalent connection of the membrane protein to the two DNA handles.

    The method may offer a long lifetime for single-molecule linkage; however, it does not represent a significant technological advancement. These reactions are commonly used in the field of single-molecule manipulation studies. The use of multiple tags including biotin and digoxygenin to enhance the connection's mechanical stability has already been explored in previous DNA mechanics studies by multiple research labs. Additionally, conducting single-molecule manipulation experiments on a single DNA or protein tether for an extended period of time (hours or even days) has been documented by several research groups.

  5. eLife

    Reviewer #3 (Public Review):

    The authors describe a method to tether proteins via DNA linkers in magnetic tweezers and apply it to a model membrane protein. The main novelty appears to be the use of DBCO click chemistry to covalently couple to the magnetic bead, which creates stable tethers for which the authors report up to >1000 force-extension cycles. Novel and stable attachment strategies are indeed important for force spectroscopy measurements, in particular for membrane proteins that are harder and therefore less studied in this regard than soluble proteins, and recording >1000 stretch and release cycles is an impressive achievement. Unfortunately, I feel that the current work falls short in some regards to exploring the full potential of the method, or at least does not provide sufficient information to fully assess the performance of the new method. Specific questions and points of attention are included below.

    - The main improvement appears to be the more stable and robust tethering approach, compared to previous methods. However, the stability is hard to evaluate from the data provided. The much more common way to test stability in the tweezers is to report lifetimes at constant force(s). Also, there are actually previous methods that report on covalent attachment, even working using DBCO. These papers should be compared.

    - The authors use the attachment to the surface via two biotin-traptavidin linkages. How does the stability of this (double) bond compare to using a single biotin? Engineered streptavidin versions have been studied previously in the magnetic tweezers, again reporting lifetimes under constant force, which appears to be a relevant point of comparison.

    - Very long measurements of protein unfolding and refolding have been reported previously. Here, too, a comparison would be relevant. In light of this previous work, the statement in the abstract "However, the weak molecular tethers used in the tweezers limit a long time, repetitive mechanical manipulation because of their force-induced bond breakage" seems a little dubious. I do not doubt that there is a need for new and better attachment chemistries, but I think it is important to be clear about what has been done already.

    - Page 5, line 99: If the PEG layer prevents any sticking of beads, how do the authors attach reference beads, which are typically used in magnetic tweezers to subtract drift?

    - Figure 3 left me somewhat puzzled. It appears to suggest that the "no detergent/lipid" condition actually works best, since it provides functional "single-molecule conjugation" for two different DBCO concentrations and two different DNA handles, unlike any other condition. But how can you have a membrane protein without any detergent or lipid? This seems hard to believe.
    Figure 3 also seems to imply that the bicelle conditions never work. The schematic in Figure 1 is then fairly misleading since it implies that bicelles also work.

    - When it comes to investigating the unfolding and refolding of scTMHC2, it would be nice to see some traces also at a constant force. As the authors state themselves: magnetic tweezers have the advantage that they "enable constant low-force measurements" (page 8, line 189). Why not use this advantage?
    In particular, I would be curious to see constant force traces in the "helix coil transition zone". Can steps in the unfolding landscape be identified? Are there intermediates?

    - Speaking of loading rates and forces: How were the forces calibrated? This seems to not be discussed. And how were constant loading rates achieved? In Figure 4 it is stated that experiments are performed at "different pulling speeds". How is this possible? In AFM (and OT) one controls position and measures force. In MT, however, you set the force and the bead position is not directly controlled, so how is a given pulling speed ensured?
    It appears to me that the numbers indicated in Figures 4A and B are actually the speeds at which the magnets are moved. This is not "pulling speed" as it is usually defined in the AFM and OT literature. Even more confusing, moving the magnets at a constant speed, would NOT correspond to a constant loading rate (which seems to be suggested in Figure 4A), given that the relationship between magnet positions and force is non-linear (in fact, it is approximately exponential in the configuration shown schematically in Figure 1).

    - Finally, when it comes to the analysis of errors, I am again puzzled. For the M270 beads used in this work, the bead-to-bead variation in force is about 10%. However, it will be constant for a given bead throughout the experiment. I would expect the apparent unfolding force to exhibit fluctuations from cycle to cycle for a given bead (due to its intrinsically stochastic nature), but also some systematic trends in a bead-to-bead comparison since the actual force will be different (by 10% standard deviation) for different beads. Unfortunately, the authors average this effect away, by averaging over beads for each cycle (Figure 4). To me, it makes much more sense to average over the 1000 cycles for each bead and then compare. Not surprisingly, they find a larger error "with bead size error" than without it (Figure 5A). However, this information could likely be used (and the error corrected), if they would only first analyze the beads separately.
    What is the physical explanation of the first fast and then slow decay of the error (Figure 5B)? I would have expected the error for a given bead after N pulling cycles to decrease as 1/sqrt(N) since each cycle gives an independent measurement. Has this been tested?

  6. eLife

    Author Response:

    Reviewer #1 (Public Review):

    This manuscript features a key technical advance in single-molecular force spectroscopy. The critical advance is to employ a click chemistry (DBCO-cycloaddition) for making a stable covalent connection between a target biomacromolecule and solid support in place of conventional antigen-antibody binding. This tweak dramatically improves the mechanical stability of the pulling system such that the pulling/relaxation can be repeated up to a thousand times (the previous limit was a few hundred cycles at best). This improvement is broadly applicable to various molecular interactions and other types of single-molecule force spectroscopy allowing for more statistically reliable force measurements. Another strength of this method is that all conjugation steps are chemically orthogonal (except for Spy-catcher conjugation to the termini of a target molecule) such that the probability of side reactions could be reduced.

    The reliability of kinetic and thermodynamic parameters obtained from single-molecule force spectroscopy depends on statistics, that is, the number of pulling measurements and their distribution. By extending the number of measurements, this robust method enables fundamental/critical statistical assessment of those parameters. That is, it is an important and interesting lesson from this study that ~200 repeats can yield statistically reasonable parameters.

    The authors carried out carefully designed optimization steps and inform readers of the critical aspects of each. The merit, quality, and rigor as a method-oriented manuscript are impressive. Overall, this is an excellent study.

    We appreciate for the positive evaluation for our work. Additionally, the minor suggestions were helpful to improve our manuscript. Thank you!

    Reviewer #2 (Public Review):

    In this study, the authors have developed methods that allow for repeatedly unfolding and refolding a membrane protein using a magnetic tweezers setup. The goal is to extend the lifespan of the single-molecule construct and gather more data from the same tether under force. This is achieved through the use of a metal-free DBCO-azide click reaction that covalently attaches a DNA handle to a superparamagnetic bead, a traptavdin-dual biotin linkage that provides a strong connection between another DNA handle and the coverslip surface, and SpyTag-SpyCatcher association for covalent connection of the membrane protein to the two DNA handles.

    The method may offer a long lifetime for single-molecule linkage; however, it does not represent a significant technological advancement. These reactions are commonly used in the field of single-molecule manipulation studies. The use of multiple tags including biotin and digoxygenin to enhance the connection's mechanical stability has already been explored in previous DNA mechanics studies by multiple research labs. Additionally, conducting single-molecule manipulation experiments on a single DNA or protein tether for an extended period of time (hours or even days) has been documented by several research groups.

    One of the unique features of our work is the development of a robust single-molecule tweezer method that is applicable to membrane proteins, rather than simply making another stable system. As re-written in Introduction, it is not straightforward as we have to consider the membrane reconstitution. We believe that our work is expected to overcome the bottleneck in membrane protein studies that arises when using single-molecule tweezer methods.

    To improve the delivery of the contextual information, we revised Introduction, Results, and Discussion. The first four paragraphs in the Introduction briefly review previous tweezer methods with an improved stability and delineate where our work is placed. In the first paragraph of the Results, we also briefly discussed how and why our DBCO tethering strategy differs from previous DBCO methods. In the first paragraph of the Discussion, we compared the previous methods regarding the stability improvement.

    Additionally, the revised manuscript now includes new findings – the full dissection of structural transitions of a helical membrane protein, the observation of hidden helix-coil transitions at a constant force, and the estimation of kinetic pre-exponential factors. We believe that the new findings provide important insights into membrane protein folding, in addition to the usefulness of our method itself for membrane protein studies. We extensively edited the main text and Methods accordingly. Relevant figures are Figures 6 and 7, Figure 6–figure supplements 1–3, and Figure 7–source data 1.

    Reviewer #3 (Public Review):

    The authors describe a method to tether proteins via DNA linkers in magnetic tweezers and apply it to a model membrane protein. The main novelty appears to be the use of DBCO click chemistry to covalently couple to the magnetic bead, which creates stable tethers for which the authors report up to >1000 force-extension cycles. Novel and stable attachment strategies are indeed important for force spectroscopy measurements, in particular for membrane proteins that are harder and therefore less studied in this regard than soluble proteins, and recording >1000 stretch and release cycles is an impressive achievement. Unfortunately, I feel that the current work falls short in some regards to exploring the full potential of the method, or at least does not provide sufficient information to fully assess the performance of the new method. Specific questions and points of attention are included below.

    We appreciate for the positive evaluation. We were able to largely improve our manuscript while preparing our responses to the comments. Thank you!

    - The main improvement appears to be the more stable and robust tethering approach, compared to previous methods. However, the stability is hard to evaluate from the data provided. The much more common way to test stability in the tweezers is to report lifetimes at constant force(s). Also, there are actually previous methods that report on covalent attachment, even working using DBCO. These papers should be compared.

    As shown in Figure 4E, we evaluated the robustness of our method in a way suggested by you – the lifetime measurement at a constant force. Specifically, ~12 hours at 50 pN. Definitely, our tweezer approach established here is the most robust method for membrane protein studies. Please refer to the section “Assessing robustness of our single-molecule tweezers” in page 7 and line 31.

    We discussed the previous covalent methods for which quantitative data are presented in light of the system stability. Please refer to the first paragraph of Discussion. We also briefly discussed how and why our DBCO tethering strategy differs from previous DBCO methods, in the first paragraph of Results.

    - The authors use the attachment to the surface via two biotin-traptavidin linkages. How does the stability of this (double) bond compare to using a single biotin? Engineered streptavidin versions have been studied previously in the magnetic tweezers, again reporting lifetimes under constant force, which appears to be a relevant point of comparison.

    The papers in this comment showed that the tethering lifetimes of biotin-streptavidin variants were affected by the asymmetric bead anchoring point. However, the situation does not apply to our work as we do not anchor traptavidin to beads. Besides, the stability comparison between the single- and double-biotin systems is not the main point of our work, so we do not have the answer to the question. However, we cited the reference in the first paragraph of Discussion where we discuss the system stability.

    - Very long measurements of protein unfolding and refolding have been reported previously. Here, too, a comparison would be relevant.

    We briefly discussed the relevant previous works in the first paragraph of Discussion.

    In light of this previous work, the statement in the abstract "However, the weak molecular tethers used in the tweezers limit a long time, repetitive mechanical manipulation because of their force-induced bond breakage" seems a little dubious. I do not doubt that there is a need for new and better attachment chemistries, but I think it is important to be clear about what has been done already.

    The sentence is in Abstract, so we also had to consider the conciseness. By simply adding the phrase “used for the membrane protein studies”, we can place our work into a more proper context.

    In page 2 and line 3, “…However, the weak molecular tethers used for the membrane protein studies have limited long-time, repetitive molecular transitions due to force-induced bond breakage…”

    - Page 5, line 99: If the PEG layer prevents any sticking of beads, how do the authors attach reference beads, which are typically used in magnetic tweezers to subtract drift?

    The PEG layer consists of biotin-PEG and methyl-PEG at a 1:27.5 molar ratio. As the reference beads are coated with streptavidin, they are attached to the PEG layer by the regular biotin-streptavidin interaction. In page 19 and line 7, you can refer to “…The polystyrene beads are attached to the PEG surface via biotin-streptavidin interaction. The beads are used as reference beads for the correction of microscope stage drifts…”

    - Figure 3 left me somewhat puzzled. It appears to suggest that the "no detergent/lipid" condition actually works best, since it provides functional "single-molecule conjugation" for two different DBCO concentrations and two different DNA handles, unlike any other condition. But how can you have a membrane protein without any detergent or lipid? This seems hard to believe.

    We explained the raised point in page 6 and line 18,

    “…Indeed, the best condition was in the absence of any detergents or lipids (Figure 3; no detergents/lipids only during the conjugation step). This situation is possible because membrane proteins are sparsely tethered to the chamber surface, which kept them from aggregating. However, not using detergents or lipids means that the membrane proteins are definitely deformed from their native folds. Therefore, we sought an optimal solubilization condition for membrane proteins during the DBCO-azide conjugation step...”

    Figure 3 also seems to imply that the bicelle conditions never work. The schematic in Figure 1 is then fairly misleading since it implies that bicelles also work.

    The buffer conditions shown in Figure 3 are those ONLY during the DBCO-azide conjugation step. In this step, the bicelle conditions did not work. Therefore, after the conjugation in 0.5% DDM, the buffer was exchanged with a bicelle solution. This process is shown in Figure 2 and the finally assembled system is depicted in Figure 1.

    To clarify this point, we put a note “Buffer conditions only during the DBCO-azide conjugation step” just above the buffer conditions in Figure 3. You can also find for the relevant exchange step in page 6 and line 31, “…Following a 1 h incubation of the beads in the single-molecule chamber at 25°C, unconjugated beads were washed, and the detergent micelles were exchanged with bicelles to reconstitute the lipid bilayer environment for membrane proteins…”

    - When it comes to investigating the unfolding and refolding of scTMHC2, it would be nice to see some traces also at a constant force. As the authors state themselves: magnetic tweezers have the advantage that they "enable constant low-force measurements" (page 8, line 189). Why not use this advantage?
    In particular, I would be curious to see constant force traces in the "helix coil transition zone". Can steps in the unfolding landscape be identified? Are there intermediates?

    Yes, please refer to Figure 6. We were able to dissect three distinct transitions from the fully unstructured state to the native state, including the helix-coil transitions. We also reconstructed the folding energy landscape using a deconvolution method.

    Please refer to the pertinent sections in the main text, which are titled “Structural transitions and folding energy landscape over extended time scales” and “Mechanistic dissection of folding transitions”.

    - Speaking of loading rates and forces: How were the forces calibrated? This seems to not be discussed.

    We wrote an additional section in Methods titled “Instrumentation of single-molecule magnetic tweezers”, where we discuss the force calibration. For the actual force calibration data, please see Figure 4–figure supplement 1A.

    In page 20 and line 10, “…The mechanical force applied to a bead-tethered molecule was calibrated as a function of the magnet position using the formula F = k_B_T∙L/_δx_2 derived from the inverted pendulum model96, where F is the applied force, _k_B is the Boltzmann constant, T is the absolute temperature, L is the extension, and _δx_2 is the magnitude of lateral fluctuations…”

    And how were constant loading rates achieved? In Figure 4 it is stated that experiments are performed at "different pulling speeds". How is this possible? In AFM (and OT) one controls position and measures force. In MT, however, you set the force and the bead position is not directly controlled, so how is a given pulling speed ensured?
    It appears to me that the numbers indicated in Figures 4A and B are actually the speeds at which the magnets are moved. This is not "pulling speed" as it is usually defined in the AFM and OT literature. Even more confusing, moving the magnets at a constant speed, would NOT correspond to a constant loading rate (which seems to be suggested in Figure 4A), given that the relationship between magnet positions and force is non-linear (in fact, it is approximately exponential in the configuration shown schematically in Figure 1).

    You are correct, so we simply modified the “pulling speed” to “magnet speed” in the figure caption. The loading rates provided in the figure (with the notation <>) were average loading rates in 1–50 pN to provide rough estimates. We actually specified it in the caption as “average force-loading rate”. However, this can be misleading at a glance, so we just deleted all the loading-rate values in the figure and caption.

    - Finally, when it comes to the analysis of errors, I am again puzzled. For the M270 beads used in this work, the bead-to-bead variation in force is about 10%. However, it will be constant for a given bead throughout the experiment. I would expect the apparent unfolding force to exhibit fluctuations from cycle to cycle for a given bead (due to its intrinsically stochastic nature), but also some systematic trends in a bead-to-bead comparison since the actual force will be different (by 10% standard deviation) for different beads. Unfortunately, the authors average this effect away, by averaging over beads for each cycle (Figure 4). To me, it makes much more sense to average over the 1000 cycles for each bead and then compare. Not surprisingly, they find a larger error "with bead size error" than without it (Figure 5A). However, this information could likely be used (and the error corrected), if they would only first analyze the beads separately.

    We might be wrong, but there seems to be a misunderstanding. First, we added Figure 5–figure supplement 1 where you can see individual traces. As expected, the levels of unfolding forces/sizes appear consistent during the progress of pulling cycles. Second, the advantage of averaging for different beads is that you can effectively remove the bead size effect. This “averaging-out” is the key strategy in our kinetic analysis. Based on the error estimation, if you average the values of kinetic parameters obtained from different beads, you can then estimate them with reasonably small errors despite the bead size variations. This becomes more evident after initial hundreds of pulling cycles. The errors for 200 and 1000 cycles are of only ~1% difference, indicating that you do not need to blindly run the pulling cycles. These results are based on the “averaging-out” strategy, which is the merit of our analysis. For more details, please see the section in the main text titled “Assessing statistical reliability of pulling-cycle experiments”, where relevant figures, figure supplements, and Method sections are referred.

    What is the physical explanation of the first fast and then slow decay of the error (Figure 5B)? I would have expected the error for a given bead after N pulling cycles to decrease as 1/sqrt(N) since each cycle gives an independent measurement. Has this been tested?

    If the sampling was from one population (here, unfolding probability profile), the error would follow a 1/√n decay as expected for the standard error. In our analysis, however, we estimated the expected “mean” errors, regardless of detailed shapes of the unfolding probability profiles. To this end, we sampled the data from different possible profiles (shown in Figure 5–figure supplement 5). We then averaged all the error plots to obtain the plot of the mean errors during progress of pulling cycles (black curve in Figure 5D). In this case, the plot does not have to follow the standard error curve represented by the factor 1/√n.

    We tested this by fitting with the model function of y = A/√n, for various lower limit of N = 10, 30, 50, 100, 300, and 500 in the regression analysis (Figure 5–figure supplement 6). The results of the reduced chi-square (χ2) used for a goodness-of-fit test (χ2 = 1 for the best fit) indicates that the two-term exponential model (χ2 = 1.60) shows a better fit than the reciprocal square root model (χ2 = 2.30–6.01). The regression model adopted in our analysis is a phenomenological model that more properly describes the error decay curve. The trend of the first fast and then slow decay is not unusual because it is also expected for the reciprocal square root model – the plot 1/√n decays fast and then slowly, too (Figure 5–figure supplement 6).

  7. Version published to 10.1101/2023.01.15.524119v1 on bioRxiv