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The next frontier: Quantitative Biochemistry in living cells Alf Honigmann, and André Nadler Biochemistry, Just Accepted Manuscript • DOI: 10.1021/acs.biochem.7b01060 • Publication Date (Web): 04 Dec 2017 Downloaded from http://pubs.acs.org on December 13, 2017
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Biochemistry
The next frontier: Quantitative Biochemistry in living cells Alf Honigmann1,* and André Nadler1,* 1
Max Planck Institute of Molecular Cell Biology and Genetics, Pfotenhauerstraße 108, 01307
Dresden, Germany *
Correspondence to André Nadler (
[email protected]), phone: +49 351 210-2970
or Alf Honigmann (
[email protected]), +49 351 210-2969
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Abstract
Researchers striving to convert biology into an exact science foremost rely on structural biology and biochemical reconstitution approaches to obtain quantitative data. However, cell biological research is moving at an ever-accelerating speed into areas where these approaches loose much of their edge. Intrinsically unstructured proteins and biochemical interaction networks composed of interchangeable, multivalent and unspecific interactions pose unique challenges to quantitative biology, as do processes that occur in discrete cellular microenvironments. Here we argue that a conceptual change in our way of conducting biochemical experiments is required to take on these new challenges. We propose that reconstitution of cellular processes in vitro should be much more focused on mimicking the cellular environment in vivo – an approach that requires detailed knowledge about the material properties of cellular compartments, essentially requiring a material science of the cell. In a similar vein, we suggest that quantitative biochemical experiments in vitro should be accompanied by corresponding experiments in vivo, as many newly relevant cellular processes are highly context-dependent. In essence, this constitutes a call for chemical biologists to convert their discipline from a proof-of-principle science to an area that could rightfully be called quantitative biochemistry in living cells. In this essay, we discuss novel techniques and experimental strategies with regard to their potential to fulfill such ambitious aims.
Introduction
How does the vast number of molecular species and reactions within cells give rise to the functional structures of a living cell? This old and fundamental question is still compelling. Curiously, it sometimes seems that the more molecular details are discovered the more difficult it
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becomes to answer this question. Our ability to observe cellular and subcellular structures and dynamics with high spatiotemporal resolution microscopy and spectroscopy techniques has undergone major advances. Consequently, our catalogue of molecular structures, metabolic reaction networks, signaling networks and gene expression patterns is growing every day. However, while we have become increasingly proficient at characterizing structures and dynamics both at the molecular and the cellular scale, we are still struggling to mechanistically explain how both regimes are connected. From the point of view of biochemists, we should ask ourselves what is holding us back to develop a mechanistic understanding of the intermediate or meso-scale regime, where molecular details are put into context and where cellular function starts to emerge? Is our reductionist approach, the reconstitution of structure and function with a minimal number of components, able to provide answers about how cellular processes selforganize in the context of highly crowded, complex and out-of-equilibrium environments? While we cannot answer this question here, we can think about the conceptual and methodological problems that need to be solved in order to make progress in this direction. In this essay, we will first shed light on a number of the conceptual problems when studying mesoscale processes in vitro. We will then discuss how quantitative biochemistry may be done directly in living cells to overcome some of the limitations of in-vitro experiments currently observed.
1. Conceptual considerations
The standard biochemical approach to understand complex biological processes is to conduct reconstitution experiments. The biomolecules in question are isolated, introduced into a suitable environment (aqueous solution or model membrane) and investigated with different methods that provide structural and dynamic information. The results are usually quantitative and allow the
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determination of binding affinities, rate constants and finally mechanistic structure function relations. This approach has been successfully used to dissect how proteins perform biochemical reactions, e.g. how enzymes catalyze reactions or how molecular motors use ATP to perform mechanical work. However, because reconstitution requires that components have to be isolated and purified from their natural environment, it is limited to proteins that are robust enough to keep their structure and function in different environments (high affinity interactions). This is one reason why for example many membrane proteins or large protein complexes are difficult to reconstitute, as their interactions are context dependent and often feature low affinities. Nonetheless, within these limitations, we have arrived at a mechanistic understanding of many different cellular sub-processes. But how single sub-processes are organized in space and time into higher level processes, for example endo/exocytosis cascades, membrane signaling pathways or a cytokinetic ring formation, has remained largely unclear. To advance biochemistry to this next level, we have to address the question how distinct biochemical reactions (subprocesses) are coupled in space and time to produce cascades of reactions with deterministic and robust outcomes.
Two major aspects have to be considered when we compare processes in living cells to reconstituted reactions in vitro: First, both the cytoplasm and the membranes of living cells are far removed from chemical equilibrium. This is readily apparent by the spontaneous, ATPindependent reorganization that both the cytosol and cellular membranes undergo when isolated. Non-equilibrium distribution means that local concentrations differ significantly across cellular space (gradients, compartments), with direct consequences for the occurrence of low-affinity interactions, which might occur in one environment but could be suppressed in another. Second, in the cellular context the concentration and diversity of macromolecules is very high and the
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distribution of components is often heterogeneous. Together, crowding of diverse molecules in an out of equilibrium situation has significant consequences for the occurrence, magnitude and lifetime of molecular interactions as compared to idealized, diluted solutions. Our current reconstitution approaches largely focus on high-affinity (lock and key) structures and interactions which are relatively insensitive to the molecular composition of their environment or its overall thermodynamic state (easy to reconstitute). In this case, the underlying free energy landscape features a small number of steep minima representing the configurational states of the assembled complex (Fig. 1B left panel). On the contrary, there are many interactions in the cell such as lipid-lipid interactions, many lipid-protein interactions and presumably also interactions between unstructured proteins that are largely non-specific or low-affinity. This does not necessarily mean that they are weak in absolute terms (measured as free energy change), but that they are interchangeable. The corresponding free energy landscape features a large number of minima that are separated by small activation energy barriers (Fig. 1B right panel). In such a case, the physico-chemical environment becomes exceedingly important. Competitive displacement of binding partners by other molecules becomes possible, residence times change and so do the overall properties of the interaction network. Therefore, the spatiotemporal control of low-affinity interaction in living cells is most likely an essential part of how meso-scale structures are assembled and how reaction cascades are regulated.
One relatively well understood example is T-cell signaling via the T-cell receptor complex. During cell signaling, information needs to be transmitted over the cell membrane to cause a cellular response. This is achieved by the formation of signaling clusters, which contain multiple activated receptors and scaffolding molecules that transiently recruit and therefore bring together kinases and other signaling proteins. The whole cluster is stabilized by actin-cortex remodeling
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and is finally inactivated by endocytosis. To understand which mechanisms actually drive the formation and termination of these signaling clusters at the cell membrane, some of its intermediate steps have been carefully studied in vivo and reconstituted in vitro
1–3
. Without
going into the molecular details here, one of the key features of the system is that formation of the signaling complex results from multivalent low affinity interactions of scaffolding proteins that are triggered by ligand induced phosphorylation of the receptor and its interacting proteins. Phosphorylation triggers a local phase transition of the system into a condensed liquid state, where components are highly concentrated but turn over quickly. Due to the high concentration of binding sites interaction partners are continuously recruited from the cytoplasm to the signaling domain and can be activated by phosphorylation. Taken together, during the process of signaling specific local molecular environments transiently self-organize in response to a defined molecular signal. These environments then facilitate interactions that do not occur in the resting state. Remarkably, each step in the cascade of the whole process seems to facilitate the formation of the next, enabling a directed progression of reactions producing a deterministic and robust outcome. Directionality necessitates that the free energy change of the entire process is negative (Fig1 C). However, the energy required for such directed processes stems not only from localized ATP hydrolysis but likely also from relaxation of out-of-equilibrium molecular distributions. This particular feature makes such processes difficult to reconstitute, as the maintenance of the initial steady state requires constant energy consumption. We chose T-cell signaling as a positive example, because it is one of the rare processes for which reconstitution of some key structures, dynamics and functions has been achieved. However, it is likely that the lessons learned from this example provide useful concepts to understand many other cellular processes.
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Indeed, most processes involving cellular membranes and non-membrane compartments which form through liquid-liquid phase separation4 are not well understood on the molecular level. In the case of biological membranes, there has been no shortage of models trying to define the connection between spatial organization of membrane components (proteins and lipids) on the mesoscale and cellular functions6–8. While not intending to argue on the merits of one or the other of the initially proposed models, it is clear that none of them have been broadly accepted as consensus models8 and they do not provide a basis for explaining directed processes at cellular membranes. An example might illustrate this point: We understand for instance clearly how the clathrin lattice is formed during clathrin mediated endocytosis, and can reconstitute it properly9, but we do not understand the molecular mechanisms that govern the temporal order of recruitment of many other effector proteins or the formation of the exclusion zone around the budding vesicle10. We simply know that both are essential for the process.
From the previous paragraphs, it follows that in vitro reconstitution of higher level mesoscale processes is becoming increasing difficult (but not impossible), because here both composition (components) and its configuration in space (compartmentalization) have to be controlled. Especially the latter is so far rarely taken care of. In our opinion, this imposes a major limitation for the classical in-vitro biochemistry reconstitution approach. We believe that a solution to this dilemma requires: (i) Development of strategies to quantitatively characterize macromolecular assemblies on the mesoscale with regard to their molecular organization principles and material properties directly in cells. (ii) This will help to reconstitute reactions in compartmentalized environments in vitro and test sufficiency of components. (iii) Development of concepts and models how nonspecific or context-dependent, often multivalent molecular interactions control and organize cellular processes. In the following paragraphs of this article, we outline how this
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challenge can be addressed, which experimental capabilities have to be developed and which techniques have the potential to be instrumental in this quest.
Figure 1: A) Three examples for the different interaction regimes that govern the assembly of bio-molecular structures. (1) Protein machineries assemble via high affinity interactions into well-defined structures (as shown for the proteasome, a DNA-Helicase and an actin filament). (2) Non-membrane bound compartments assemble via multivalent interactions between proteins and occasionally RNA oligonucleotides and low affinity interactions of intrinsically disordered protein domains. (3) Structure formation in and at lipid membranes crucially relies on lowaffinity lipid-lipid and lipid-protein interactions. (B) Schematic difference of the free energy landscapes of the two different regimes. Protein machineries feature steep energy minima representing stable configurational states of the complex (left panel). Non-membrane bound
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compartments and lipid membranes typically exhibit only small activation energy barriers between the local minima representing many different configurational states, rendering these assemblies highly dynamic and context dependent (right panel). C) Trajectory of a cellular process. The steady state of the compartment is “poised” for the initiation of a directed, deterministic cascade of transitions by virtue of a steady state far removed from chemical equilibrium which is maintained through constant ATP consumption. A specific trigger allows the cascade to start and each intermediate features a particular molecular organization that might enhance distinct low affinity interactions while preventing others.
1. Material science of the cell
One of the underlying problems in understanding biochemical reaction networks in cells is our limited ability to characterize the local environments in which the reactions are actually taking place. It has long been known that the cytoplasm as well as cellular membranes are very crowded environments, with a dense packing of proteins, RNA, lipids, metabolites and salts. While the general effect of simple macro-molecular crowding on biochemical reactions has been well established11, it is becoming more and more clear that complex crowded environments facilitate segregation of specific components into local compartments and environments4. In membranes sub-compartmentalization induced by collective lipid-lipid and lipid-protein segregation mechanisms has been extensively studied in vitro and in vivo over the last 20 years12. While many of the fundamental questions in membrane field are still open, the recent discovery of nonmembrane bound compartments in the nucleoplasm and the cytoplasm shows that low affinity interactions and collective effects not only play a crucial role in membranes but are key in the organization of cellular space in general4,13. Clearly, this discovery opens exciting new
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perspectives to understand many cellular processes. But what tools do we actually have to study the underlying molecular interactions of these structures? Lessons from the membrane organization field have shown that attempts to simply reconstitute or isolate membrane microenvironments often produces results that are difficult to compare to cellular data8. This is at least partially to blame on choosing components to mimic cellular complexity in reconstitution experiments based on pragmatism rather than on actual knowledge of the respective cellular micro-environment. To move forward and solve some of the discrepancies between in vitro and in vivo experiments we think it is essential to strive for a quantitative characterization not only of the specific protein of interest but also of its local environment in the cellular context. Key parameters that should be determined inside and outside the respective structure or microenvironment are concentrations of the main protein and/or lipid components, RNA/DNA content, metabolites (ATP, NADH etc.), salt concentrations (Ca2+, Mg2+), pH and viscosity. This is of course not a trivial task, but we believe it is absolutely necessary to make sense of experimental data. The following sections will give a brief overview of current methods for quantitative description of cellular environments and interactions.
1.1. Fluorescence and its limitations for quantification of sub-cellular environments
The method of choice for live imaging of cellular processes is fluorescence microscopy, as it combines high contrast with molecular specificity in a non-invasive way. Fluorescence tagging of endogenous cellular molecules is straightforward for proteins using genetically encoded tags, which are either auto-fluorescent14 or can be coupled to organic dyes under live cell conditions15. Using current genome engineering tools such as CRISPR/Cas9
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16
, proteins of interest can be
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tagged and expressed at endogenous concentrations (provided the tag is not interfering with protein localization and function). Obtaining a quantitative measurement from a fluorescence imaging experiment requires careful calibrations to convert the detected signal into molecular numbers17. In case the molecules of interest are mobile, fluorescent fluctuation spectroscopy methods (e.g. FCS and its derivatives18) are the tool of choice. Traditional FCS provides average molecular concentration and mobility in a single femto-liter volume (confocal detection spot). Recent advancements such as Number and Brightness and SPIM-FCS allow mapping concentration and mobility across section of the cell in 2D19–21. In principle these new tools are ideally suited to detect and quantify heterogeneous molecular environments and interactions based on concentration and mobility of the respective molecules in space. To further deduce the underlying causes for heterogeneity and non-Brownian motion of molecules FCS measurements have to performed on different spatial scales22,23. This allows discriminating active processes, confined diffusion and molecular trapping from standard Brownian motion. Since FCS is based on measuring stochastic fluctuations of molecules in space and time it should be possible to determine on which time and spatial scales cellular molecules behave purely stochastic and on which scale non-random and robust patterns emerge. While FCS provides time and ensemble averages (in the tens of second range), single molecule tracking methods directly reveal the complexity of molecular movements and interactions in space and time24, albeit with the limitation of operating only at very low concentrations.
Unfortunately, for non-protein bio-molecules, fluorescence labelling is not straightforward. One option is to chemically attach an organic dye to the molecule of interest. In addition to the problem that these molecules have to be introduced into cells from the outside, modification of small molecules with relatively large aromatic moieties such as dyes changes the physico-
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chemical properties and therefore their molecular identity. A prominent example are fluorescent lipid analogues, which can be used to measure general membrane properties but have only very limited value to understand the function of specific lipids25. The current workaround to this problem is to use engineered fluorescent proteins that serve as sensors for specific molecules. For example, sensors to probe pH, ion concentrations or certain signaling lipids are available26,27. While the number of available sensors is steadily increasing, so far only a very limited number of molecules can be probed. Additionally, absolute quantification of concentrations based on the biosensor signals is difficult and it requires laborious calibration of the sensors and the microscopes. Hence, most studies report only relative changes. Finally, the sensor itself is typically overexpressed and therefore care has to be taken not to interfere with the cellular processes. An alternative to protein based sensors for probing cellular environments are organic environmentally sensitive fluorescent dyes. Different small probes are available for measuring viscosity, hydrophobicity, salinity and temperature28. Unfortunately, most of the available probes respond to more than property simultaneously, resulting in difficult analysis of data stemming from complex environments. In addition, such probes typically do not localize to a defined compartment, so that the obtained signal is an average stemming from multiple different environments. A promising strategy in this regard is the development of environmentally sensitive groups that are targeted to particular cellular compartments through specific chemical groups29,30. The final problem when using either fluorescent protein sensors or environmentally sensitive dyes is the diffraction limited resolution of conventional microscopy. Electron micrographs of cellular cross-sections show a dense packing of cellular organelles (e.g. ER, mitochondria etc.), which are intermingled by cytoplasm. The resolution of the best confocal microscopes is 600nm in the Z direction and 200nm in the XY plane. Therefore, conventional
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imaging unavoidably averages over organelles and cytoplasm. So far, super-resolution methods, which offer significant higher resolution, require specific dyes which in many cases are not compatible with the current biosensors31. However, some promising combinations with superresolution microscopy have been reported32.
Taken together, fluorescent tools to quantify cellular micro-environments are available. However, so far only a small number of molecules and parameters can be probed. Using the tools to create quantitative maps of sub-cellular environments has so far been hampered by the low resolution of conventional microscopes.
1.2. Alternatives to fluorescence microscopy
Fluorescence microscopy has the advantage of molecular specificity. The downside is that only a very limited number of molecular species can be detected simultaneously (typically up to three) and often with low spatial resolution. (Cryo)Electron microscopy, on the other hand, has the opposite problem. It offers very high spatial resolution and all molecules in the sample contribute to the image. Recent advances in microscope design and sample handling enable imaging of cellular sections up to a thickness of 100 nm with a resolution and contrast that reveals the ultrastructure of large protein complexes and membranes in their native environment33. The dream to create spatial maps of all the major cellular machineries within cells is coming into reach. Unfortunately, small molecules and proteins that are not assembled into well-defined structures are not yet visible in cryo-EM. Combing cryo-EM with fluorescence microscopy to bridge this gap is a very promising direction34. Another alternative to characterize cellular space in a non-
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invasive manner are label-free imaging techniques, such as coherent anti-Stokes Raman (CARS), multiphoton microscopy, and confocal Raman microscopy. Raman microscopy in principle allows to detect molecular species based on their vibrational spectra with diffraction-limited resolution. However, current implementations have two major limitations. First, the contrast of the measured spectra in the complex cellular environment is very low. Second, extracting quantitative molecular information from complex spectra is so far only possible for a very limited number of molecules. While these limitations are not fundamental it will require significant progress until these methods will help us to reveal biochemistry in living cells. To detect molecular species with the highest sensitivity and precision mass spectrometry has become the method of choice. Because conventional mass spectrometry is an in vitro technique, analyzing molecular composition of sub-cellular environments relies on cellular fractionation and purification protocols, which are often non-quantitative. A technically challenging but promising direction is the use of secondary ion mass spectrometry in an imaging configuration (nanoSIMS)35. A primary ion beam is used to shoot of secondary ions from the surface of the sample of interest (e.g. fixed cells). The secondary ions are analyzed in mass spectrometry mode, which allows to map the elemental, isotopic, and with limitations also the molecular composition with a lateral resolution of up to 50 nm. Pioneering studies have shown proof of principle in fixed cells and tissues and even combination with fluorescence microscopy36,37. Unfortunately, nanoSIMS devices are still rare and the technique has to still mature into a useful and quantitative tool for biochemistry and cell biology. Taken together, tools to characterize the molecular composition, ultra-structure and dynamics of the sub-cellular space are becoming increasingly powerful. We believe that it is time to systematically apply these tools in
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combination with biochemical experiments to shine light on the self-organization of complex environments and its coupling to biochemical reactions.
2. Biochemistry in living cells – Methodological challenges
Reconstitution is by definition a reductionist approach – the attempt to find the minimal set of components that still faithfully recreates the cellular process. The key question is whether this reconstitution as a standalone-strategy can be utilized to study interactions that crucially depend on complex cellular microenvironments. We believe that this is not the case as even a reconstitution matrix chosen to perfectly mimic the properties of a chosen cellular environment will always fall short of an exact replication, due to what might be called the “unknown unknowns”: We will likely never be able to monitor the exact molecular composition of a given cellular compartment in space and time. This basic fact means that there will always be a discrepancy between molecular interactions in vitro and in vivo for certain molecular interactions. Far from being solely detrimental, this discrepancy could constitute a discovery mechanism if we were able to combine quantitative in vitro data with directly comparable measurements in vivo. The purpose of reconstitution in this situation would not to recreate the cellular process but to serve as a measure for gauging the influence of the cellular environment on the investigated process. However, this scenario has yet to be realized. In essence, it calls for the ability to measure binding affinities between biomolecules and reaction rates in living cells in absolute terms, essentially arguing for the development of biochemistry in living cells. Being able to accurately measure absolute molecular concentrations, densities and partition coefficients as described above is only a starting point. Conducting actual biochemical experiments requires additional experimental advances, particularly when it comes to small molecules, lipids and
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nucleotides. Our current repertoire is clearly insufficient to allow describing a cellular phenomenon in the quantitative thermodynamic and kinetic terms of classical biochemistry, in Kd and Km values, in rate constants, activation energies and free energy changes. The most fundamental experimental capacity for such quantitative thermodynamic and kinetic measurements is the ability to vary the concentration of crucial components on fast time scales. In living cells, inducing protein concentration changes through knockdown or overexpression of a target gene is probably the most commonly used perturbation. However, the cell will invariably adapt its overall protein composition in response to genetic perturbations, thus causing a systematic error in all quantitative measurements. Aggravating these difficulties, attempts to genetically alter the levels of non-protein biomolecules are fraught with difficulties and rarely yield satisfying results. Consequently, conducting quantitative biochemical experiments in living cells hinges on technological advances in two main areas: The ability to induce rapid, exactly defined concentration changes of biomolecules on millisecond to second timescales and a rapid expansion of techniques to quantitatively monitor the intracellular distribution and concentration of non-protein biomolecules in live-cell fluorescence microscopy experiments as discussed above.
2.1.Chemical and physical approaches for controlling biomolecule concentration and cellular material properties
Changing the intracellular concentration of biomolecules on rapid timescales without resorting to pharmacological or genetic approaches has long been the domain of Chemical Biology. A number of approaches have been developed over the last decades. The oldest and still most straightforward is probably photo-induced release of biomolecules from so called photo-caged
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precursor compounds383940. UV-induced removal of a photo-cleavable protection group positioned on a functional group of crucial biological importance during a live-cell fluorescence microscopy or patch-clamp experiment triggers a step-wise concentration increase of a distinct molecular species and thus creates an opportunity to investigate its biological role. A significant number of such compounds have been reported in the literature, and their utilization has become a staple of cell biological (primarily neurological) studies40. However, to be truly useful in actual quantitative in vivo biochemical experiments, a number of obstacles have to be removed. First, while uncaging does in fact trigger step-wise concentration increases, the exact amount of photoreleased biomolecule is typically not known, thus prohibiting quantitative analysis in principle41. Apart from utilizing standardized biosensors for measuring the concentration of the released biomolecules, methods for quantifying the intracellular concentration of the caged parent molecule as well as measuring the efficiency of the uncaging photoreaction will have to be improved41. Second, caged compounds do not typically localize to distinct cellular compartments. A typical uncaging experiment thus elevates the concentration of the target molecule indiscriminately in the entire cell. In principle, there are two possibilities to address this issue. On the one hand, localized (preferably two-photon) uncaging allows for the generation of intracellular concentration gradients. However, localized uncaging approaches likely always result in photo-release in multiple organelles at the same time, largely due to the crowded intracellular space, as discussed above. On the other hand, it is possible to pre-localize caged components to individual organelles, as demonstrated for the plasma membrane41, the nucleus42 and mitochondria43 (Figure 2A). A key challenge in the field will thus be to expand the available repertoire of organelle-targeting caging groups and to make thoroughly characterized compounds available to the cell biological community.
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Photo-switchable small molecules offer a complementary method to control biomolecule function with light44. The most commonly used scaffolds are based on azobenzenes (Figure XX)44. Photo-switches can be used in a number of ways, but are most prominently applied as ligands for cell-surface receptors45, as inhibitors of intracellular proteins46 or as mimics of second messengers 47. The major advantage compared to caged compounds lies in the reversibility of the induced effects. A good example is provided by recently reported photo-switchable DAGs
47
,
which enable to induce oscillatory recruitment of DAG binding proteins to cellular membranes (Figure 2B). The major disadvantage of photo-switchable second messenger mimics lies in the fact that incorporation of diazobenzene structures represents a significant chemical alteration compared to the natural parent molecule. The major challenges for further development towards quantitative applications of photo-switchable tools are comparable to the obstacles faces by researchers developing new caged compounds. Photoreactions need to be quantified inside living cells, the effective intracellular concentrations of photo-switchable actuators must be readily accessible as well as affinities of both photo-isomers for target proteins. Going forward, caged small molecules will likely remain the method of choice for studying endogenous small molecules in their natural context, whereas chemical photo-switches offer unique possibilities for investigating temporal patterns of protein activity, especially when genetic approaches are not accessible.
In addition to small molecule uncaging and photo-switching approaches, the use of chemical dimerizers as tools for inducing rapid intracellular concentration changes has grown in popularity 4849
. Chemically induced dimerization (CID) approaches rely on triggering a protein-protein
interaction through external application of a small molecule (Figure 2C). Localizing one of the protein binding partners at a specific cellular compartment thus allows to elevate the
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concentration of the other protein in that compartment in a second to minute time regime based on the membrane permeability of the utilized chemical dimerizer. Furthermore, by recruiting enzymes to particular cellular localizations, it is also possible to locally alter the levels of signaling lipids and other small molecules. CID has a number of intrinsic advantages that make it particularly suited for quantitative biochemistry in vivo. If the recruited protein is directly involved in the investigated process, its local concentration can be determined by straightforward fluorescence microscopy approaches. Furthermore, CID in principle allows to compare the kinetic properties of enzymes in different microenvironments. Next, due to the usual prelocalization of one of the binding partners, discrimination between different cellular compartments is usually not an issue. Potential drawbacks are to be seen in inherently slower onset kinetics (in the second to minute range), which lowers the applicability of this technology in fast cellular processes. The ongoing development of reversible and photo-caged chemical dimerizers might solve this issue in the future48. A more fundamental challenge is the fact that the usefulness of the CID technology as a quantitative method is currently largely limited to proteins. Recruiting an enzyme to a particular cellular compartment changes the concentration of its substrates and products, but quantifying the exact magnitude of these changes is often impossible. Progress in this area will likely come from further development of quantitative biosensors.
In contrast to small molecule-centered approaches as discussed above, optogenetic tools rely on light-sensitive proteins to induce rapid changes of cellular environments5051. By now, a number of optogenetic modules exist, which rely on different molecular principles (Figure 2E)52. Lightsensitive bacterial rhodopsins serve as tools to modulate membrane potential and intracellular ion concentrations. Variants of these proteins serve as engineered G-protein coupled receptors,
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effectively enabling the light-controlled activation of cellular signaling cascades. LOV domain and Cry2 based tools have been central for the development of protein homo- and heterodimerization systems and the development of allosterically activated kinases, effectively constituting a complementary strategy to CID52. Due to rapid progress in recent years, optogentic tools now enable the modulation of numerous cellular processes, including membrane potential changes, cellular signaling cascades, gene expression, nuclear import and protein secretion. Optogenetic approaches on the one hand feature faster activation kinetics than CID, but microscopic characterization of the induced effects is usually more limited in scope as a significant part of the visible spectrum is required for inducing the conformational changes. Both approaches offer similar advantages and drawbacks for controlling small molecule and protein concentrations, as optogenetic control of intracellular small molecule levels is similarly dependent on enzymatic transformations.
In addition to optogenetic tools that target the activity of a single protein or the interaction between well-defined binding partners, a number of tools have been reported that allow to modulate cellular properties in a more general manner. Prominent examples include the variation of cellular contractility by controlling RhoA activity53, control of organelle positioning with the IM-LARIAT system54 and light-triggered phase transitions in the cytoplasm55. Very recently, a new technique termed FLUCS has been reported that allows to generate cytosolic flows without relying on light-sensitive proteins57. Essentially, laser radiation is used to generate localized temperature waves of low magnitude (2-3 °C) but high frequency. Due to a complex interplay between thermal expansion phenomena and viscosity changes, localized flows emerge that can be finely tuned by variation of excitation patterns. The proposed method represents an elegant approach to physically manipulate living cells without relying on protein function (Figure 2D).
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From the current point of view, it seems likely that photochemical uncaging will be the method of choice for triggering small molecule concentration changes, whereas CID, photoswitches or optogenetic approaches seem to be preferable for rapid changes of local protein concentrations, control of protein function and more generally modulation of alterations of cellular environments. In all cases, the most significant benefits for the field of in-vivo biochemistry are to be expected if further technological development is focused on generating tools that allow for precise modulation and quantification of cellular parameters rather than extending the number of existing proof-of-principle approaches.
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Figure 2: Experimental approaches for rapid and targeted perturbations of cellular processes A) Plasma-membrane specific photo-caged compounds allow to elevate second messenger levels allow for subcellular modulation of second messenger levels41. B) Photoswitchable diacylglycerol derivative enable reversible recruitment of Protein Kinase C47. C) Small-molecule induced dimerization enables re-localization of cellular protein pools to specific intracellular sites48. D) FLUCS: Cytoplasmic flows are generated through high-frequency temperature waves [57]. E) Most common principles of optogenetic systems52.
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3. Conclusion
Biochemistry, at its heart, is the attempt to understand how the transformations and interactions of biomolecules are translated into cellular function. Traditionally, reconstitution of biochemical reactions and interactions in vitro has been the primary source of quantitative data. However, if we want to understand how the biochemical reactions are controlled in space and time, we have to go one step further and include the environment of the respective system. This poses a challenge to the reductionist biochemical approach of reconstitution, because it is becoming increasingly clear that the specificity of molecular interactions and the structural complexity of the cellular environment are highly interdependent. In this essay, we argued that an over-reliance on in vitro approaches will be insufficient to properly characterize biological processes in such environments. Reconstitution approaches should always be accompanied by corresponding measurements in living cells and the results of the two approaches need to be compared. In this scenario, a reductionist approach would not be an attempt to recreate but rather a discovery mechanism. Divergences between quantitative measurements in vivo and in vitro would point towards not-yet-discovered components of the investigated network. It will take more than a well-meaning opinion article to make this vision a reality. Currently, the vast majority of microscopy-based cell biological studies report only relative data in individually chosen metrics – publications by the authors are no exception. If we aim at routinely performing quantitative live cell experiments, we need a cultural shift with regard to analysing microscopy data as well as development and implementation of new experimental approaches. Instead of reporting instrument dependent fluorescent intensities, existing methods to quantify protein concentrations
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in live cell experiments should find more widespread application. A key problem that is still largely unsolved is the microscopic visualization of non-protein biomolecules. The generation of analogues and conjugates by the chemical biology community clearly has not sufficed and better methods for visualizing endogenous small molecules, lipids and nucleotides are needed. Ultimately, this will likely mean a rapid expansion of the number of available biosensors both for soluble species and membrane lipids. Next, we need experimental avenues for conducting quantitative kinetic measurements in living cells. A number of promising techniques such as photochemical uncaging, chemical dimerizers and optogenetic protein dimerization exist. The key challenge will be to improve these techniques to a point where defined and scalable alteration of the concentrations of distinct molecular species in individual cellular compartments becomes possible. Ideally, quantitative results from in vivo experiments should be readily comparable between laboratories. To this end, it would be immensely helpful if a central database would exist where all published binding affinities, Km values, partition coefficients and protein concentrations would be deposited. Ultimately, solving the current limitations in understanding the molecular biology of the cell will require a holistic approach combining cell biological, biochemical, chemical biological and biophysical approaches.
Funding Information
The research of A. N. has received funding from the European Research Council (ERC) under the European Union’s Horizon 2020 research and innovation program (grant agreement no GA 758334 ASYMMEM) and the Deutsche Forschungsgemeinschaft (DFG) as member of the TRR83 consortium. The research of A. H. has received funding from the Deutsche Forschungsgemeinschaft (DFG) as member of the TRR83 consortium.
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[57]
Mittasch, M., Groß, P., Nestler, M., Fritsch, A. W., Iserman, C., Kar, M., Munder, M.,
Voigt, A., Alberti, S., Grill, S. W., Kreysing, M. “Non-invasive perturbations of intracellular flow reveal physical principles of cell organization”, in press.
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For table of contents use only
Title: The next frontier: Quantitative Biochemistry in living cells Authors: Alf Honigmann1,* and André Nadler1,*
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