Chemistry & Biology
Previews Light Moves Mountains in the Cell Sohum Mehta1 and Jin Zhang1,2,* 1Department
of Pharmacology and Molecular Sciences, The Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA Solomon H. Snyder Department of Neuroscience and Department of Oncology, The Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA *Correspondence: [email protected] http://dx.doi.org/10.1016/j.chembiol.2015.05.002 2The
In this issue of Chemistry & Biology, Duan et al. (2015) report the use of a light-inducible protein-protein interaction system to dynamically control the movement of intracellular organelles with spatial and temporal precision in living cells. A quick glance in any biology text book reveals that intracellular organelles are routinely depicted as adopting a highly stereotypical arrangement, with a central nucleus surrounded by the ER and neighbored by the Golgi, while mitochondria, lysosomes, and other organelles are often dispersed peripherally in the cytoplasm. In reality, however, organelles are highly dynamic and motile entities whose movements are controlled by motor proteins such as kinesin and dynein (Barlan et al., 2013), which mediate the transport of diverse molecular cargoes, including entire organelles, along microtubules. Alterations in the distribution of organelles are essential for numerous cellular processes, including cell division, polarized morphogenesis, and migration (Bornens, 2008; Gundersen and Worman, 2013). Organelles also change their positions in response to various external stimuli— mitochondria move toward the nucleus in response to a lack of oxygen, or hypoxia (Al-Mehdi et al., 2012), while lysosomes alter their distribution based on the availability of nutrients (Korolchuk et al., 2011) —and decades of research indicate that organelle positioning can have a profound impact on a variety of biological processes. Yet the functional consequences of organelle positioning are not always clear; elucidating these effects requires tools to directly manipulate how organelles are distributed within living cells. To this end, Duan et al. (2015) describe a straightforward method for dynamically controlling organelle localization based on the rapidly expanding field of optogenetics. Optogenetics entails the use of genetic techniques to make living cells responsive to light in order to turn on or off specific events within these cells (Deisseroth,
2011). Optical methods are ideal for selectively perturbing cellular events with high spatial and temporal resolution because light can be focused to submicron levels and applied with millisecond timing. The use of light to control cellular processes is possible because many organisms natively respond to light and express proteins whose activities are naturally light sensitive. By studying these organisms, scientists have identified a number of protein domains that can be inserted into proteins of interest to render their functions directly sensitive to light. As a rule, these light-controlled proteins contain chemical moieties (e.g., chromophores) that absorb light at specific wavelengths; the ensuing photo-chemical reaction then leads to conformational changes that alter protein function. Arabidopsis thaliana cryptochrome 2 (CRY2), for example, contains a flavin cofactor that absorbs blue light and induces a conformational change that promotes binding to the transcription factor CIB1. The use of this and similar light-sensitive protein domains has allowed scientists to exert optical control over protein-protein interactions and even various enzyme activities (Zhang and Cui, 2015). Yet, as optogenetics has matured, it has also shifted from focusing solely on the activities of macromolecules to manipulating larger cellular processes. As if to help underscore this shift—from perturbing proteins to moving mountains—the present study by Duan et al. (2015) uses the CRY/CIB system to control the movement of organelles by tethering them to specific motor proteins. Specifically, the authors localized CRY2 to different organelle membranes and fused CIB1 to either KIF5A (e.g., kinesin) or to the N-terminal domain of the
dynein-binding BICD protein (BICDN), which complements a recently described system that uses a modified light-oxygenvoltage (LOV) domain-PDZ interaction to control organelle movement (van Bergeijk et al., 2015). Kinesin and dynein both ‘‘walk’’ along microtubules, which typically radiate outward from the vicinity of the nucleus. Kinesin moves toward microtubule plus-ends, located near the cell edge, and was thus used to move organelles away from the nucleus. Conversely, dynein moves toward microtubule minus-ends and was used to direct organelles inward, toward the nucleus. Thus, the authors tethered CRY2, tagged with the red fluorescent protein mCherry, to the transmembrane domain of the mitochondrially localized Miro1 protein (MiroTM). When this was co-expressed with GFP-BICDN-CIB1 in COS-7 (monkey) cells, the authors were able to successfully induce the redistribution of mitochondria to the perinuclear region following blue-light exposure. Similarly, by co-expressing CRY2-mCherry-MiroTM with KIF5A-GFP-CIB1, the authors were able to use blue light to increase mitochondrial localization at the extreme periphery of cells. This design was extended to other organelles, namely, peroxisomes and lysosomes, which the authors directed toward the nucleus or the cell edge depending on tethering to dynein or kinesin, respectively. The authors also performed an important control by labeling microtubules with YFP and confirming that these light-recruited motors are indeed moving along microtubules. This also allowed the authors to quantify the speed of light-induced peroxisome movement: both kinesin- and dynein-mediated peroxisome transport matched rates described in the literature (Kural et al.,
Chemistry & Biology 22, May 21, 2015 ª2015 Elsevier Ltd All rights reserved 569
Chemistry & Biology
Previews 2005). Interestingly, the authors noted that only a subset of microtubules was used for organelle transport, with many microtubules carrying multiple peroxisomes. Two essential goals of optogenetic approaches are to control cellular events reversibly and with a high degree of spatial precision. To highlight the reversibility of their system, Duan and coworkers demonstrated that CIB1-GFPMiroTM and CRY2-mCherry dimers could be rapidly and repeatedly assembled and disassembled in response to alternating periods of blue light and darkness, with no apparent loss of dimerization efficiency over time. Furthermore, mitochondria that were induced to relocalize following blue-light exposure returned to their original distribution in the dark. The authors similarly demonstrated the spatial selectivity of their optogenetic system, through which they were able to induce the redistribution of mitochondria toward the cell edge only within a defined cellular region without altering mitochondrial
positioning elsewhere in the cell. In addition, a key property of the CRY/CIB dimerization system is its activation by very low light intensities, which the authors highlight here by showing that COS-7 cells expressing CRY2-mCherry-MiroTM with either KIF5A-GFP-CIB1 or BICDN-GFPCIB1 could be exposed to low levels of blue light for extended periods of time without any cell death. These tools could therefore provide a fresh perspective on old biological questions, such as how organelle positioning within specific cellular regions affects processes such as polarized cell growth, as well as how organelle migration influences developmental processes. Even more exciting, however, is the prospect that these new molecular tools will also force us to pose new questions that we had not considered previously.
V.V., Alexeyev, M.F., and Gillespie, M.N. (2012). Sci. Signal. 5, ra47. Barlan, K., Rossow, M.J., and Gelfand, V.I. (2013). Curr. Opin. Cell Biol. 25, 483–488. Bornens, M. (2008). Nat. Rev. Mol. Cell Biol. 9, 874–886. Deisseroth, K. (2011). Nat. Methods 8, 26–29. Duan, L., Che, D., Zhang, K., Ong, Q., Guo, S., and Cui, B. (2015). Chem. Biol. 22, this issue, 671–682. Gundersen, G.G., and Worman, H.J. (2013). Cell 152, 1376–1389. Korolchuk, V.I., Saiki, S., Lichtenberg, M., Siddiqi, F.H., Roberts, E.A., Imarisio, S., Jahreiss, L., Sarkar, S., Futter, M., Menzies, F.M., et al. (2011). Nat. Cell Biol. 13, 453–460. Kural, C., Kim, H., Syed, S., Goshima, G., Gelfand, V.I., and Selvin, P.R. (2005). Science 308, 1469– 1472.
REFERENCES
van Bergeijk, P., Adrian, M., Hoogenraad, C.C., and Kapitein, L.C. (2015). Nature 518, 111–114.
Al-Mehdi, A.-B., Pastukh, V.M., Swiger, B.M., Reed, D.J., Patel, M.R., Bardwell, G.C., Pastukh,
Zhang, K., and Cui, B. (2015). Trends Biotechnol. 33, 92–100.
570 Chemistry & Biology 22, May 21, 2015 ª2015 Elsevier Ltd All rights reserved
Previews Light Moves Mountains in the Cell Sohum Mehta1 and Jin Zhang1,2,* 1Department
of Pharmacology and Molecular Sciences, The Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA Solomon H. Snyder Department of Neuroscience and Department of Oncology, The Johns Hopkins University School of Medicine, Baltimore, MD 21205, USA *Correspondence: [email protected] http://dx.doi.org/10.1016/j.chembiol.2015.05.002 2The
In this issue of Chemistry & Biology, Duan et al. (2015) report the use of a light-inducible protein-protein interaction system to dynamically control the movement of intracellular organelles with spatial and temporal precision in living cells. A quick glance in any biology text book reveals that intracellular organelles are routinely depicted as adopting a highly stereotypical arrangement, with a central nucleus surrounded by the ER and neighbored by the Golgi, while mitochondria, lysosomes, and other organelles are often dispersed peripherally in the cytoplasm. In reality, however, organelles are highly dynamic and motile entities whose movements are controlled by motor proteins such as kinesin and dynein (Barlan et al., 2013), which mediate the transport of diverse molecular cargoes, including entire organelles, along microtubules. Alterations in the distribution of organelles are essential for numerous cellular processes, including cell division, polarized morphogenesis, and migration (Bornens, 2008; Gundersen and Worman, 2013). Organelles also change their positions in response to various external stimuli— mitochondria move toward the nucleus in response to a lack of oxygen, or hypoxia (Al-Mehdi et al., 2012), while lysosomes alter their distribution based on the availability of nutrients (Korolchuk et al., 2011) —and decades of research indicate that organelle positioning can have a profound impact on a variety of biological processes. Yet the functional consequences of organelle positioning are not always clear; elucidating these effects requires tools to directly manipulate how organelles are distributed within living cells. To this end, Duan et al. (2015) describe a straightforward method for dynamically controlling organelle localization based on the rapidly expanding field of optogenetics. Optogenetics entails the use of genetic techniques to make living cells responsive to light in order to turn on or off specific events within these cells (Deisseroth,
2011). Optical methods are ideal for selectively perturbing cellular events with high spatial and temporal resolution because light can be focused to submicron levels and applied with millisecond timing. The use of light to control cellular processes is possible because many organisms natively respond to light and express proteins whose activities are naturally light sensitive. By studying these organisms, scientists have identified a number of protein domains that can be inserted into proteins of interest to render their functions directly sensitive to light. As a rule, these light-controlled proteins contain chemical moieties (e.g., chromophores) that absorb light at specific wavelengths; the ensuing photo-chemical reaction then leads to conformational changes that alter protein function. Arabidopsis thaliana cryptochrome 2 (CRY2), for example, contains a flavin cofactor that absorbs blue light and induces a conformational change that promotes binding to the transcription factor CIB1. The use of this and similar light-sensitive protein domains has allowed scientists to exert optical control over protein-protein interactions and even various enzyme activities (Zhang and Cui, 2015). Yet, as optogenetics has matured, it has also shifted from focusing solely on the activities of macromolecules to manipulating larger cellular processes. As if to help underscore this shift—from perturbing proteins to moving mountains—the present study by Duan et al. (2015) uses the CRY/CIB system to control the movement of organelles by tethering them to specific motor proteins. Specifically, the authors localized CRY2 to different organelle membranes and fused CIB1 to either KIF5A (e.g., kinesin) or to the N-terminal domain of the
dynein-binding BICD protein (BICDN), which complements a recently described system that uses a modified light-oxygenvoltage (LOV) domain-PDZ interaction to control organelle movement (van Bergeijk et al., 2015). Kinesin and dynein both ‘‘walk’’ along microtubules, which typically radiate outward from the vicinity of the nucleus. Kinesin moves toward microtubule plus-ends, located near the cell edge, and was thus used to move organelles away from the nucleus. Conversely, dynein moves toward microtubule minus-ends and was used to direct organelles inward, toward the nucleus. Thus, the authors tethered CRY2, tagged with the red fluorescent protein mCherry, to the transmembrane domain of the mitochondrially localized Miro1 protein (MiroTM). When this was co-expressed with GFP-BICDN-CIB1 in COS-7 (monkey) cells, the authors were able to successfully induce the redistribution of mitochondria to the perinuclear region following blue-light exposure. Similarly, by co-expressing CRY2-mCherry-MiroTM with KIF5A-GFP-CIB1, the authors were able to use blue light to increase mitochondrial localization at the extreme periphery of cells. This design was extended to other organelles, namely, peroxisomes and lysosomes, which the authors directed toward the nucleus or the cell edge depending on tethering to dynein or kinesin, respectively. The authors also performed an important control by labeling microtubules with YFP and confirming that these light-recruited motors are indeed moving along microtubules. This also allowed the authors to quantify the speed of light-induced peroxisome movement: both kinesin- and dynein-mediated peroxisome transport matched rates described in the literature (Kural et al.,
Chemistry & Biology 22, May 21, 2015 ª2015 Elsevier Ltd All rights reserved 569
Chemistry & Biology
Previews 2005). Interestingly, the authors noted that only a subset of microtubules was used for organelle transport, with many microtubules carrying multiple peroxisomes. Two essential goals of optogenetic approaches are to control cellular events reversibly and with a high degree of spatial precision. To highlight the reversibility of their system, Duan and coworkers demonstrated that CIB1-GFPMiroTM and CRY2-mCherry dimers could be rapidly and repeatedly assembled and disassembled in response to alternating periods of blue light and darkness, with no apparent loss of dimerization efficiency over time. Furthermore, mitochondria that were induced to relocalize following blue-light exposure returned to their original distribution in the dark. The authors similarly demonstrated the spatial selectivity of their optogenetic system, through which they were able to induce the redistribution of mitochondria toward the cell edge only within a defined cellular region without altering mitochondrial
positioning elsewhere in the cell. In addition, a key property of the CRY/CIB dimerization system is its activation by very low light intensities, which the authors highlight here by showing that COS-7 cells expressing CRY2-mCherry-MiroTM with either KIF5A-GFP-CIB1 or BICDN-GFPCIB1 could be exposed to low levels of blue light for extended periods of time without any cell death. These tools could therefore provide a fresh perspective on old biological questions, such as how organelle positioning within specific cellular regions affects processes such as polarized cell growth, as well as how organelle migration influences developmental processes. Even more exciting, however, is the prospect that these new molecular tools will also force us to pose new questions that we had not considered previously.
V.V., Alexeyev, M.F., and Gillespie, M.N. (2012). Sci. Signal. 5, ra47. Barlan, K., Rossow, M.J., and Gelfand, V.I. (2013). Curr. Opin. Cell Biol. 25, 483–488. Bornens, M. (2008). Nat. Rev. Mol. Cell Biol. 9, 874–886. Deisseroth, K. (2011). Nat. Methods 8, 26–29. Duan, L., Che, D., Zhang, K., Ong, Q., Guo, S., and Cui, B. (2015). Chem. Biol. 22, this issue, 671–682. Gundersen, G.G., and Worman, H.J. (2013). Cell 152, 1376–1389. Korolchuk, V.I., Saiki, S., Lichtenberg, M., Siddiqi, F.H., Roberts, E.A., Imarisio, S., Jahreiss, L., Sarkar, S., Futter, M., Menzies, F.M., et al. (2011). Nat. Cell Biol. 13, 453–460. Kural, C., Kim, H., Syed, S., Goshima, G., Gelfand, V.I., and Selvin, P.R. (2005). Science 308, 1469– 1472.
REFERENCES
van Bergeijk, P., Adrian, M., Hoogenraad, C.C., and Kapitein, L.C. (2015). Nature 518, 111–114.
Al-Mehdi, A.-B., Pastukh, V.M., Swiger, B.M., Reed, D.J., Patel, M.R., Bardwell, G.C., Pastukh,
Zhang, K., and Cui, B. (2015). Trends Biotechnol. 33, 92–100.
570 Chemistry & Biology 22, May 21, 2015 ª2015 Elsevier Ltd All rights reserved
