COPI/COPII proteins and kinesins/dyneins

COPI/COPII proteins and kinesins/dyneins

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I am considering the transport of protein from ER to Golgi, and have read that this involves the COPII protein coat. I have also read that this is a form of anterograde transport, and elsewhere that kinesins are responsible for anterograde transport as they move from the negative to positive (outside) poles of the microtubules.

Thus it seemed to me that the kjnesins carry the COPII vesicles carrying proteins from ER to Golgi.

However a Google search of the terms 'kinesins' and 'COPII' together did not seem to yield anything about kinesins carrying COPII.

On the other hand in this article it links COPII with dynsctjn, which I have not heard of before.

Does someone know the link between COPI and COPII transport, and/or retrograde and anterograde transport, with the microtubule motor proteins kinesin and dynein? Where do dynectins come into this? Do they bind the vesicle to the dynein or are they a different type of motor protein? If it is the first case, then why is this transport termed anterograde if it does not use kinesins? Are the terms anterograde and retrograde not directly related to kinesin/dyneins use?

Interesting question. The term anterograde refers to movement in the forward direction. In the context of vesicular trafficking, anterograde refers to (1) movement from the site of protein synthesis in the rough endoplasmic reticulum (RER) towards the Golgi and then (2) movement from the Golgi towards the final destination in the cell.

Both processes rely on microtubules for transport. Microtubules are polar and radiate from the microtubule organizing center (MTOC) with their (+) ends directed towards the periphery of the cell:

There are two broad classes of molecular motors that facilitate directed movement along microtubules. Kinesins are generally (+)-end directed motors whereas dyneins are (-)-end directed:

These images should already give you a hint. Your question may have arisen due to the assumption that the ER is centrally located near the MTOC with the Golgi somewhat peripheral. This is incorrect. In reality, the Golgi complex is actively clustered near the MTOC by movement on microtubules. Furthermore, the ER extends along the microtubule network towards the (+)-end periphery. Taken together, it is evident that microtubule (-)-ends are located near the Golgi and transport to it from any part of the cell, including the RER, requires the (-)-end directed motor dynein. Subsequent anterograde transport from the Golgi to a specific destination in the cell, or through the secretory pathway, requires the (+)-end directed motor kinesin. Retrograde transport from the Golgi back to the ER would also use kinesin:

The article cited in the question (which I also reference in this answer), mentions that COPII vesicles are coupled to microtubules via dynactin. Dynactin is a protein complex that is used to both activate and adapt cargo to dynein (ie COPII vesicles are coupled to microtubules via a dynein/dynactin complex):

COPI/COPII proteins and kinesins/dyneins - Biology

A tendency in cell biology is to divide and conquer. For example, decades of painstaking work have led to an understanding of endoplasmic reticulum (ER) and Golgi structure, dynamics, and transport. In parallel, cytoskeletal researchers have revealed a fantastic diversity of structure and cellular function in both actin and microtubules. Increasingly, these areas overlap, necessitating an understanding of both organelle and cytoskeletal biology. This review addresses connections between the actin/microtubule cytoskeletons and organelles in animal cells, focusing on three key areas: ER structure and function ER-to-Golgi transport and Golgi structure and function. Making these connections has been challenging for several reasons: the small sizes and dynamic characteristics of some components the fact that organelle-specific cytoskeletal elements can easily be obscured by more abundant cytoskeletal structures and the difficulties in imaging membranes and cytoskeleton simultaneously, especially at the ultrastructural level. One major concept is that the cytoskeleton is frequently used to generate force for membrane movement, with two potential consequences: translocation of the organelle, or deformation of the organelle membrane. While initially discussing issues common to metazoan cells in general, we subsequently highlight specific features of neurons, since these highly polarized cells present unique challenges for organellar distribution and dynamics.


Transport carriers operating between early compartments in the mammalian secretory pathway have to travel long distances in the cell by mostly relying on the microtubule network and its associated motor proteins. Although anterograde transport from the endoplasmic reticulum (ER) to the Golgi complex is mediated by cytoplasmic dynein 1, 2, the identity of the motor(s) mediating transport in the retrograde direction is presently unclear. Some studies have suggested that the heterotrimeric kinesin-2 complex plays a role in transport between the ER and the Golgi 3, 4. Here, we have examined kinesin-2 function by using an RNA-interference approach to downregulate the expression of KAP3, the nonmotor subunit of kinesin-2, in HeLa cells. KAP3 silencing results in the fragmentation of the Golgi apparatus and a change in the steady-state localization of the KDEL-receptor (KDEL-R). Using specific transport assays, we show that the rate of anterograde secretory traffic is unaffected in these cells but that KDEL-R-dependent retrograde transport is strongly abrogated. Our data strongly support a role for kinesin-2 in the KDEL-R-/COPI-dependent retrograde transport pathway from the Golgi complex to the ER.

Present address: Institució Catalana de Recerca I Estudis Avançats and Centre de Regulacio Genomica, Dr. Aiguader 88, 08003 Barcelona, Spain.

The trafficking protein Tmed2/p24β1 is required for morphogenesis of the mouse embryo and placenta

During vesicular transport between the endoplasmic reticulum and the Golgi, members of the TMED/p24 protein family form hetero-oligomeric complexes that facilitate protein-cargo recognition as well as vesicle budding. In addition, they regulate each other's level of expression. Despite analyses of TMED/p24 protein distribution in mammalian cells, yeast, and C. elegans, little is known about the role of this family in vertebrate embryogenesis. We report the presence of a single point mutation in Tmed2/p24β1 in a mutant mouse line, 99J, identified in an ENU mutagenesis screen for recessive developmental abnormalities. This mutation does not affect Tmed2/p24β1 mRNA levels but results in loss of TMED2/p24β1 protein. Prior to death at mid-gestation, 99J homozygous mutant embryos exhibit developmental delay, abnormal rostral–caudal elongation, randomized heart looping, and absence of the labyrinth layer of the placenta. We find that Tmed2/p24β1 is normally expressed in tissues showing morphological defects in 99J mutant embryos and that these affected tissues lack the TMED2/p24β1 oligomerization partners, TMED7/p24γ3 and TMED10/p24δ1. Our data reveal a requirement for TMED2/p24β1 protein in the morphogenesis of the mouse embryo and placenta.

Molecular motors drive the transport of vesicles and organelles within the cell. Traditionally, these transport processes have been considered separately from membrane trafficking events, such as regulated budding and fusion. However, recent progress has revealed mechanistic links that integrate these processes within the cell. Rab proteins, which function as key regulators of intracellular trafficking, have now been shown to recruit specific motors to organelle membranes. Rab-independent recruitment of motors by adaptor or scaffolding proteins is also a key mechanism. Once recruited to vesicles and organelles, these motors can then drive directed transport this directed transport could in turn affect the efficiency of trafficking events. Here, we discuss this coordinated regulation of trafficking and transport, which provides a powerful mechanism for temporal and spatial control of cellular dynamics.

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The zebrafish toolbox – microscopes, genetics and drugs

Zebrafish overview

Zebrafish produce large numbers of externally fertilized eggs, which rapidly develop into transparent embryos and progress to free-swimming feeding larvae within 5 days post fertilization (dpf). Gastrulation is complete within hours of fertilization and by 1 dpf, embryonic axes are established and neural development is underway. An exquisite map of cell division and movement during these early developmental events has been achieved by tracking individually labeled nuclei using advanced microscopy (Keller et al., 2008 Schmid et al., 2013). By the end of 2 dpf, organogenesis is underway throughout the embryo and, by 5 dpf, most organs carry out specialized functions. Although zebrafish have traditionally been used to study embryogenesis, areas such as behavior, pathology, infectious diseases and drug screening are actively investigated. Here, we focus on zebrafish tools used to study cell biology in vivo, with the aim to understand development and disease.

Labeling subcellular structures in zebrafish

High-resolution studies of embryonic development and disease models require analysis of the behaviors of individual cells, subcellular structures and proteins in the context of an intact tissue or organism. Fluorescent proteins have revolutionized cell biology. However, the relative small size and dynamic nature of subcellular components present challenges for imaging. This is exacerbated by the cellular complexity of whole animals. Thus, in contrast to the elegant studies describing organelle dynamics in cultured cells or yeast, relatively little is known about these processes in vertebrates.

Developing a zebrafish toolkit that labels, tracks and measures intracellular structures comparable to that available in mammals is a work in progress. A recent and concerted effort by the zebrafish community and several companies has improved the library of antibodies recognizing zebrafish proteins, which are cataloged in the zebrafish model organism database ( However, specific antibodies for many structures remain unavailable, and antibody staining does not allow live imaging, a strength of the zebrafish system. Developing transgenics (Kawakami, 2005 Kwan et al., 2007) and vital dyes provides a good alternative. Well-characterized fluorescent markers relevant to topics discussed here are listed in Table 1 and illustrated in Fig. 1F. Fig. 1 shows two examples of such transgenics: Tg(bact:GalT-GFP) fish express a portion of Galactose-1-phosphate uridyl transferase (GALT) fused to GFP under the semi-ubiquitous β-actin promoter (Gerhart et al., 2012) to enable imaging of the trans-Golgi in the developing embryo using both a low-resolution fluorescence stereomicroscope (Fig. 1A–C) and confocal imaging (Fig. 1D). The second, Tg(ef1a:dclk-GFP) line (Tran et al., 2012), labels microtubules with GFP and, when used in combination with the vesicle marker clathrin fused to dsRed (M.S. and M.M., unpublished data Fig. 1E), it provides a real-time view of microtubule-based vesicular transport.

Fluorescent protein markers of subcellular structures in zebrafish. (A–C) Live 2 dpf Tg(bact:GalT-GFP) embryo highlighting the trans-Golgi complex. The boxed region in B is magnified in the C. (D) Confocal projection through a cryosection of the liver from 5 dpf Tg(bact:GalT-GFP) larva. Nuclei are stained with DAPI (gray). (E) Muscle in a live 1 dpf Tg(ef1a:dclk-GFP) embryo with GFP-tagged microtubule-associated protein (DCK) and vesicles marked with clathrin-DS-Red (magenta). (F) Zebrafish fluorescent-protein-tagged organelle markers that can be expressed with transgenics. PM, plasma membrane LE, late endosomes EE, early endosomes RE, recycling endosomes.

Fluorescent protein markers of subcellular structures in zebrafish. (A–C) Live 2 dpf Tg(bact:GalT-GFP) embryo highlighting the trans-Golgi complex. The boxed region in B is magnified in the C. (D) Confocal projection through a cryosection of the liver from 5 dpf Tg(bact:GalT-GFP) larva. Nuclei are stained with DAPI (gray). (E) Muscle in a live 1 dpf Tg(ef1a:dclk-GFP) embryo with GFP-tagged microtubule-associated protein (DCK) and vesicles marked with clathrin-DS-Red (magenta). (F) Zebrafish fluorescent-protein-tagged organelle markers that can be expressed with transgenics. PM, plasma membrane LE, late endosomes EE, early endosomes RE, recycling endosomes.

This regulates GFP translation to reflect endogenous Chop levels and monitor the ER stress response. GFP is found in neural tissues during ER and thermal stress.

Fluorescent vital dyes also allow for imaging subcellular structures in vivo. BODIPY, Alizarin Red, quantum dots and dyes, such as MitoTracker or LysoTracker, are versatile counterstains to visualize dynamic cellular processes in live fish (Cooper et al., 2005 He et al., 2009 Köster and Fraser, 2006 Rieger et al., 2005 Zhang et al., 2008). In combination, transgenics and dyes provide a microscopic view of intracellular processes and a macroscopic view of how disrupting basic cellular processes interrupt development or cause disease.


Many zebrafish imaging studies are carried out using microscopes that are available to most researchers, whereas an in-depth analysis of dynamic movements or morphometric analyses of subcellular structures requires the advanced microscopy techniques limited to researchers who have specialized training and access to facilities with these capabilities.

Low-resolution epifluorescent and stereomicroscopes can be used to detect gross changes in localization or intensity of many markers, and standard confocal microscopy can image cells that are close to the surface of live embryos but imaging deeper cells requires histological sections. Unprecedented views of how cells move individually and in concert in a developing embryo have been provided using light-sheet microscopy to track individual nuclei labeled with fluorescently-tagged histones (Huisken and Stainier, 2009 Keller et al., 2008). Selective plane illumination microscopy has provided a three-dimensional view of endodermal cell movement in older live embryos (Schmid et al., 2013). Multiphoton imaging has been used to investigate how neuronal circuits assemble during retinal development by tracking dynamic changes in cell structure and connectivity (Williams et al., 2013), and super-resolution microscopy has been used to image a single receptor molecule uncovering the role for receptor clustering in the antiviral immune response (Gabor et al., 2013). Many more spectacular zebrafish images are being generated as imaging technologies continue to improve.


The zebrafish genome has been sequenced and annotated, and most zebrafish genes are highly conserved in mammals, with a zebrafish ortholog identified for ∼70% of human genes (Howe et al., 2013). Transgenesis and forward genetic screens are important widely used methods in zebrafish, and recent advances in reverse genetic approaches are having a major impact on the field. Box 2 details methods of generating transgenics and targeted mutagenesis using TALENs and the Crispr/Cas systems.

Forward genetic screens allow the unbiased discovery of genes that contribute to a specific phenotype and have generated thousands of mutants in the zebrafish genome. One drawback, however, is that generating and maintaining stable lines can be laborious and another is that the genome duplication that occurred during teleost evolution (Postlethwait et al., 1998) can generate genes that act redundantly, necessitating that both are targeted before revealing a phenotype. In addition, maternal stores of mRNA and proteins can delay the emergence of mutant phenotypes until after these are exhausted, and thus mutant phenotypes often reflect the impact of gene loss on later developmental stages.

Morpholinos provide a complementary approach, whereby oligonucleotides that are injected into the fertilized egg block target mRNA translation or splicing. Although the use of morpholinos is plagued by fears of off-target effects and transient effects, translation-blocking morpholinos deplete proteins derived from both maternal and zygotic mRNA, so that the effects of target gene loss on early embryonic events can be studied. Additionally, by titrating the amount of morpholino injected, the degree of knockdown can be finely tuned. This has proved particularly useful for our work to model one type of congenital disorder of glycosylation (CDG), which, in humans is caused by a hypomorphic mutation in one of the genes required for N-linked protein glycosylation (Freeze, 2007). Homozygous null mutations of these genes are lethal in mammals (Thiel and Körner, 2011) and injecting high morpholino concentrations causes severe embryonic phenotypes and high mortality in zebrafish (Chu et al., 2013 Cline et al., 2012). However, fine-tuning of the knockdown was facilitated by injecting lower morpholino concentrations so that residual enzyme activity in the zebrafish morphants matched that measured in samples from patients, improving survival and revealing novel phenotypes (Chu et al., 2013 Cline et al., 2012). Although the breadth and speed of zebrafish genetic approaches do not match those available in invertebrates, they are accomplished at a fraction of the costs of, and with sample sizes exceeding, typical rodent experiments (see Box 1).


Zebrafish have a rich history in toxicology research, as compounds can be simply added to their water. For example, zebrafish are proving useful for alcohol research (Jang et al., 2012 Monk et al., 2013 North et al., 2010 Passeri et al., 2009 Tsedensodnom et al., 2013 Yin et al., 2012). By using transgenic zebrafish expressing a GFP-tagged secreted glycoprotein in hepatocytes (Howarth et al., 2013 Tsedensodnom et al., 2013 Xie et al., 2010), and zebrafish that express fluorescent protein markers of the hepatocyte secretory organelles and of other cells in the liver (Yin et al., 2012), the mechanisms by which alcohol and other drugs cause organ-specific and organelle-mediated toxicity can be uniquely addressed. Moreover, large-scale drug screens exploit the ease of treating zebrafish with drugs (Peterson and Macrae, 2012) and have identified compounds modulating processes ranging from metabolism (Nath et al., 2013) to sleep (Rihel and Schier, 2012).

Exocytosis in Neurons

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Synaptic vesicle exocytosis occurs in neurons of the nervous system. Nerve cells communicate by electrical or chemical (neurotransmitters) signals that are passed from one neuron to the next. Neurotransmitters are transmitted by exocytosis. They are chemical messages that are transported from nerve to nerve by synaptic vesicles. Synaptic vesicles are membranous sacs formed by endocytosis of the plasma membrane at pre-synaptic nerve terminals.

Once formed, these vesicles are filled with neurotransmitters and sent toward an area of the plasma membrane called the active zone. The synaptic vesicle awaits a signal, an influx of calcium ions brought on by an action potential, which allows the vesicle to dock at the pre-synaptic membrane. Actual fusion of the vesicle with the pre-synaptic membrane does not occur until a second influx of calcium ions occurs.

After receiving the second signal, the synaptic vesicle fuses with the pre-synaptic membrane creating a fusion pore. This pore expands as the two membranes become one and the neurotransmitters are released into the synaptic cleft (gap between the pre-synaptic and post-synaptic neurons). The neurotransmitters bind to receptors on the post-synaptic neuron. The post-synaptic neuron may either be excited or inhibited by the binding of the neurotransmitters.

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