A mitochondrial protein complex that links apoptosis and glycolysis

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The functions of retrograde transport

The best-studied function of retrograde transport is the recycling of receptors which are in charge of cargo exchange between the biosynthesis/secretory and the endocytic pathways.
Mannose 6-phosphate receptors (MPRs) in mammalian cells and Vps10 in the budding yeast Saccharomyces cerevisae capture the newly synthesized acid hydrolases at the TGN and transport them to endosomes. Upon endosomal acidification, the ligands are dissociated from their receptors (Munier-Lehmann, Mauxion et al. 1996). The hydrolases are then transferred to lysosomes, and the receptors are recycled by retrograde transport to TGN in order to enter a new functional cycle (Duncan and Kornfeld 1988; Marcusson, Horazdovsky et al. 1994). Multiligand type-I receptors such as sortilin, sorLA and sorCS, the homologues of Vps10 in mammalian cells (Mari, Bujny et al. 2008) function similarly.
The current list of proteins that cycle between endosomes and the Golgi complex includes the SNAREs (soluble N-ethylmaleimide-sensitive fusion factor attachment receptors), VAMP4 and GS15 (Tai, Lu et al. 2004; Tran, Zeng et al. 2007) which are mostly engaged in membrane fusions. Retrograde transport is also involved in development, according to recent studies. The Wnt proteins are secreted signaling molecules that play a central role in development and adult tissue homeostasis. Retrograde transport is involved in Wnt function via the trafficking of the Wnt receptor Wntless (Eaton 2008). Wntless is a conserved transmembrane protein that binds Wnt and is required for Wnt secretion. After Wntless has transported Wnt from the TGN to the plasma membrane, Wntless needs to be retrieved by retrograde transport to the TGN for a new functional cycle. Cells may also regulate retrograde transport aiming to control the range of Wnt signaling in the tissue (Yang, Lorenowicz et al. 2008).
By regulating retrograde transport, cells can balance intracellular ion homeostasis. The Menkes P-type ATPase (MNK), a copper transporter, follows retrograde transport between plasma membrane and Golgi apparatus to regulate intracellular copper levels. In cells subjected to low extracellular copper concentrations, transport of MNK from the Golgi complex is slower than retrograde retrieval from the cell membrane. MNK is consequently accumulated in the Golgi and the intracellular copper concentration is kept at normal levels. If extracellular copper concentrations are increased, the rate of MNK exocytosis is increased compared to retrograde transport so that the copper can be removed from cells (Petris and Mercer 1999). Similar function exist also in yeast for the iron transporter Fet3p/Ftr1p (Strochlic, Setty et al. 2007) whose transport is triggered by changes in iron concentration, and the mammalian glucose transporter GLUT4 which uses the hormone insulin as its trafficking signal in adipocytes and skeletal muscle cells (Shewan, van Dam et al. 2003).
Membrane-type matrix metalloproteinases (MT-MMP) are found on specialized cell surface structures, the so-called invadopodia (Poincloux, Lizarraga et al. 2009).
These proteases change their surrounding environment during angiogenesis, tissue remodeling, tumor invasion, and metastasis by cleavage of extracellular matrix components (Rowe and Weiss 2008). MT-MMP are internalized from the plasma membrane into endosomes and the TGN, and recycled back to the cell surface (Steffen, Le Dez et al. 2008). Endocytosis and recycling enables the cell to control MT-MMP activity during matrix degradation and to maintain a dynamic pool of MT-MMPs which can be used to adapt cell surface in response to physiological and pathological signals. The molecular machinery underlying retrograde transport of MT-MMP from endosomes to the TGN still needs to be identified.


Shiga Toxin

Retrograde transport is used as an entry pathway for protein toxins such as Shiga toxin, ricin, and cholera toxin. Shiga toxins-producing bacteria were first described by the Japanese microbiologist Kiyoshi Shiga in 1897. Shiga toxin producing E. coli infects more than 73.000 people every year in the US alone, and has generated annual health care costs of about 400 million dollars in 2003 (Frenzen, Drake et al. 2005). The Shiga toxins family includes Shiga toxin (STx) from of Shigella dysenteriae, and Shiga-like toxins (SLT; or verotoxins) from Escherichia coli. E. coli produces different variants of two types of SLT, Shiga-like toxin 1 (Stx1; very similar to Shiga toxin) and a Shiga-like toxin 2 family of molecules (Stx2). Shiga toxin is composed of one A-subunit, consisting of the domains A1 and A2, which is non-covalently associated with one homopentameric Shiga toxin B-subunit (STxB). STxB binds the cellular toxin receptor, the glycosphingolipid globotriaosyl ceramide (Gb3) (Figure 6).
Figure 6. The structure of shiga toxin. Shiga toxin consists of an A-subunit that is non-covalently attached to a B-subunit composed of five identical fragments. The A-subunit is cleaved by the protease furin into an enzymatically active A1-fragment and a carboxyl terminal A2-fragment, which remain linked by a disulfide bond.(Sandvig, Bergan et al. 2010)
After receptor binding, Shiga toxin is internalized to early endosomes, and then transported along retrograde transport route to the Golgi, and eventually to the ER. In the ER, the cellular ER-associated protein degradation (ERAD) machinery removes misfolded, misassembled, and/or misglycosylated proteins from the ER lumen for proteasomal degradation in the cytosol (Bernasconi and Molinari 2011). Shiga toxin A-subunit uses this ERAD machinery to escape from the ER lumen to the cytosol where it cleaves a specific adenine nucleobase from 28S RNA of the 60S subunit of the ribosome, thereby inhibiting protein synthesis.


Ricin is found in castor beans (Ricinus communis). Ricin is composed of the catalytic A-subunit and a monomeric B-subunit which functions for receptor binding. These two subunits are covalently linked together by a disulfide bond. Ricin B-subunit has two binding pockets (Sphyris, Lord et al. 1995) for galactose and N-acetylgalactoseamine residues of glycoproteins and glycolipids (Baenziger and Fiete 1979). Ricin is taken up into cells by multiple endocytosis pathways and transported to the ER by retrograde transport (Lord, Jolliffe et al. 2003).

Cholera toxin

Cholera toxin (CTx) is a protein complex secreted by the bacterium Vibrio cholera. This bacterium causes profuse watery diarrhea and vomiting in humans. Intestinal fluid loss is caused primarily by the release of cholera toxin, in addition to related enterotoxins, after adhesion of Vibrio cholera to the proximal small intestine of the host. Cholera affects 3-5 million people and causes 100,000 to 130,000 deaths a year.
The structure of Cholera toxin is similar to that of Shiga toxin: both are composed of a monomeric A-subunit and a homopentameric B-subunit. The transport route of Cholera toxin in cell is also similar to that of Shiga toxin. Instead of binding to Gb3, the B-subunit of Cholera toxin binds to the glycos phingolipid mono sialotetra hexosylgan glioside (GM1).


Besides toxins, some viruses also rely on retrograde transport. The HIV-1 envelop protein (Env) was shown to be a cargo of this pathway (Blot, Janvier et al. 2003). Depletion of retrograde transport proteins dramatically inhibits HIV replication (Lopez-Verges, Camus et al. 2006). In another study, host proteins that are required for HIV infection identified in a genome-wide RNA interference screen. This study revealed a critical role for Rab6 and its guanine nucleotide exchange factor Rgp1p in HIV-1 infection (Brass, Dykxhoorn et al. 2008). HIV immune evasion is mediated by the viral protein Nef, which induces the retrograde transport of MHC class I and costimulatory molecules (Blagoveshchenskaya, Thomas et al. 2002; Chaudhry, Das et al. 2007). The life cycle of several other viruses similarly requires retrograde transport. For example, the nonenveloped simian virus 40 (SV40) and other polyomaviruses rely on retrograde transport to the ER for their entry into the cytosolic (Spooner, Smith et al. 2006).

Other diseases

Some diseases like Alzheimer’s can also be related to retrograde transport. APP (amyloid-β precursor protein) is processed after endocytosis by the β-secretases BACE1 and BACE2, which cycle between endosomes and the TGN (He, Li et al. 2005). Subsequent cleavage by γ-secretase eventually leads to the formation of a 40-or 42-residue peptide named Aβ. Excessive levels of Aβ cause serious dysfunctions and may be a reason for the progression of Alzheimer disease (Thinakaran and Koo 2008). Other studies on Alzheimer also indicate that the dysfunction of retrograde transport can be associated with late-stage-onset of the disease (Small 2008). In neuronal cells, APP interacts with the Vps10 domain-containing sorting receptor sorLA, a cycling transport of the retrograde route (see above). Genetic studies confirmed a role of sorLA in the progression of Alzheimer’s disease (Rogaeva, Meng et al. 2007).
The Alzheimer-associated beta-secretase BACE1 interacts with Golgi-localized, gamma ear-containing, ADP ribosylation factor-binding proteins (GGA) that play a role in the sorting of cargo proteins from the TGN to endosomal compartments. It has been demonstrated that GGA1 also regulates the retrograde transport of internalized BACE1 from endosomal compartments to the Golgi through a direct interaction that is sensitive to phosphorylation (Wahle, Prager et al. 2005). While phosphorylated BACE1 is efficiently transported from endosomes to the Golgi, non-phosphorylated BACE1 enters the recycling route to the cell surface.

Retrograde transport machinery

Different elements of molecular machinery that is implicated in the retrograde transport as well as their functions are listed in Figure 7 and Table 2.
Figure 7. The location of different elements in retrograde transport (Johannes and Popoff 2008)
Table 2. Proteins involved in endosome-to-Golgi transport (Johannes and Popoff 2008)

Clathrin coats

Clathrin is a protein that plays a major role in the formation of coated vesicles. Clathrin was first isolated and named by Barbara Pearse in 1975 (Pearse 1976). Each clathrin subunit is composed of three clathrin heavy chains and three light chains that together form a three-legged structure called triskelion (Ungewickell and Branton 1981). Clathrin triskelia self-assemble into a polyhedral cage (Blank and Brodsky 1987) (Figure 8). Clathrin can be found on the plasma membrane, the TGN, and early endosomes. It plays an important role in endocytic uptake of plasma membrane receptors. However, clathrin is unable to bind directly to membranes and needs adaptor proteins (AP) such as AP-1 and AP-2 that simultaneously bind to clathrin lipids, and protein components in the membrane (Ahle and Ungewickell 1989).
EpsinR is a clathrin adaptor that mediates retrograde transport from endosomes to the TGN. Like any member of the epsin family, epsinR comprises a phosphoinositide-binding ENTH domain. Its ENTH-domain can bind efficiently to PtdIns4P, a phosphoinositide that is found on Golgi membranes and possibly on endosomes (Hirst, Motley et al. 2003). In addition, epsinR binds to AP-1. Due to these interactions with AP-1 and PtdIns4P, epsinR is localised to endosomes and the TGN.
Clathrin coats are critical for retrograde transport from endosomes to the TGN. RNAi-mediated depletion of clathrin or epsinR leads to a block of STxB in early endosomes (Saint-Pol, Yelamos et al. 2004). Similarly, retrograde transport of MPR and TGN46 also relies on epsinR. Immunogold labeling studies revealed the presence of MPR on AP-1-containing clathrin-coated vesicles and its function in MPR trafficking to the TGN (Klumperman, Hille et al. 1993).
Figure 8. The structure of clathrin and the position of clathrin triskelions in a clathrin coat (Kirchhausen 2000)

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Besides clathrin, retromer has also been described to mediate vesicle transport between early endosomes and the TGN (Seaman, McCaffery et al. 1998). Retromer is an evolutionarily conserved protein complex composed of a trimeric vps subunit, composed of Vps26, Vps29 and Vps35, and a dimer of the BAR-domain-containing sortin nexins SNX1, SNX2, SNX5, and SNX6. Double knock-outs of SNX1/SNX2 in mice and deletion of Vps35 in Drosophila melanogaster will result in embryonic lethality (Collins 2008). These studies demonstrate the importance of retromer for cell function. SNX proteins contain a SNX-phox homology (SNX-PX) domain, by which SNX are recruited onto PtdIns3P and PtdIns(3,5)P2 enriched endosomes (Teasdale, Loci et al. 2001). The combination of phosphoinositide-binding PX and BAR-domains leads the SNX proteins to be recruited to endosomal membranes and to facilitate the formation of endosomal membrane tubules.
Retromer has been demonstrated to be critical for the retrograde transport of several cargo proteins like Wntless, MPR, and Shiga toxin (Bonifacino and Hurley 2008). For several of these cargos, clathrin and clathrin-interaction proteins are also required. A recent model explains how clathrin and retromer may function in successive steps in the formation of retrograde transport intermediates (Popoff, Mardones et al. 2007). In short, clathrin would drive membrane curvature changes and the formation of retrograde tubules. Retromer would be critical for the subsequent processing of retrograde tubules. Proteins have been identified bind both, clathrin and retromer. They could explain the molecular details of the articulation between the two machineries.

Tools to study retrograde transport

The different analysis methods to study retrograde transport are listed in Table 3. The advantage and disadvantage of those methods are also shown. The currently most popular tool in cell biology is immunofuorescence to study the intracellular distribution of toxins by microscopy, followed by an analysis of their relocalization after inhibition of retrograde transport (Johannes, Tenza et al. 1997; Amessou, Popoff et al. 2006). The toxins can be labeled with fluorescent dyes, after which they can be followed without a need for immunolabeling. Furthermore, during the initial phase of their uptake into cells, the toxins are transported in a unidirectional manner. This is different from the pool of endogenous proteins that in most cases are at equilibrium between forward and backward movements. In most cases, people use only the receptor-binding parts of the toxins in cell biology studies. These usually preserve the intracellular transport characteristics of the holotoxins without being themselves toxic.
The coupling between HRP and toxins allows the observation of the cytosolic surface of organelles by electron microscopy using a technique called the whole mount. The study of endosomal coats in the retrograde route is an example for this technique (Saint-Pol, Yelamos et al. 2004). A limitation of immunofluorescence is that it is not very quantitative. STxB-based conjugates have therefore been developed to quantitatively study the different transport steps that constitute the retrograde route. Retrograde transport to the Golgi apparatus can be measured by using a specific peptide sequence that is a recognition sequence for TGN-localized sulfotransferase. When a STxB molecule that carries a tandem of this peptide sequence, termed STxB-Sulf2, reaches the TGN by retrograde transport from the plasma membrane, sulfotransferase catalyzes the transfer of inorganic sulfate from the medium onto the conserved peptide sequence. This modification can be quantitatively detected with radioactive sulfate. This technique on which one of my strategies was based will be described in detail in the next chapter.
Table 3. Tools for analysis of the retrograde route (Johannes and Popoff 2008).
In vitro assays have been developed to exploit sulfation analysis for the reconstitution of endosomes-to-TGN transport on permeabilized cells or membrane fractions (Mallard, Tang et al. 2002). In these assays, the activity of the cytosol can be directly modified with antibodies, mutant proteins or chemical inhibitors. This powerful assay is technically difficult to handle and needs a lot of training to be mastered.
Glycosylation analysis applies a similar principle as sulfation for measuring cargo arrival in the ER using ER-localized oligosaccharyl transferase and glycosylation site-tagged STxB (Fujinaga, Wolf et al. 2003).
The sulfation and glycosylation tools can also be applied to the study of retrograde transport of endogenous cargo proteins. The different sulfation or glycosylation trafficing probes are introduced onto these cargos either directly, or via antibodies. It becomes a critical issue to know whether during the time of modification, enough endogenous cargos molecules are present at the plasma membrane to generate a signal that can subsequently be detected.


The term “proteome” was used for the first time in 1995 (James 1997) to describe the protein complement of a genome. Rapidly, the name of a new discipline was coined, proteomics. In essence, proteomics is the study of protein properties (expression levels, post-translational modifications, interactions etc.) on a large scale to obtain a global, integrated view of disease processes, cellular processes and networks at the protein level.
With the dramatically increased amounts of DNA sequences in databases, it is realized that only having complete sequences of genomes is not sufficient to elucidate biological function. A cell depends on a complicated series of metabolic and regulatory pathways for its survival. There is no strict relationship between genes sequence and the cell functions. Proteomics is necessarily complementary to genomics because it focuses on the proteins that are the active elements in cells. Therefore, proteomics directly contributes to reveal cell functions.
Proteomics can be mainly divided into three areas: (1) protein characterization for large scale identification of proteins; (2) “di fferential display” proteomics for comparison of protein levels; and (3) studies of protein–protein interactions.

The functions and methods of proteomics

Identification and analysis of proteins

The most significant breakthrough in proteomics is the mass spectrometric identification of gel-separated proteins (Figure 9). The proteins to be analyzed are isolated from cell lysates by biochemical fractionation or affinity purification, often followed by gel electrophoresis. Usually, the proteins are then digested with a protease such as trypsin. The mass spectrometer is used to determine the masses of the digested peptides. Thereby, a mass map can be obtained. This mass map is then compared with predicted mass maps of proteins within the database to eventually identify the proteins.
The basic components of all mass spectrometers are the same. These are composed from an ionization source, a mass analyzer, and an ion detector. The ionization source is used to convert molecules into gas phase ions. For protein analysis, the two most common types of ionization sources are matrix-assisted laser desorption ionization (MALDI) and electrospray ionization (ESI). The development of ESI for the analysis of biological macromolecules (Kebarle and Verkerk 2009) was rewarded with the attribution of the Nobel Prize in Chemistry to John Bennett Fenn in 2002.
Figure 9. A strategy for mass spectrometric identification of proteins (Lueking, Horn et al. 1999).
In MALDI, the ions are created by using a laser to excite a crystalline mixture of analyte into the gas phase. ESI employs a potential difference between a capillary and the inlet of the mass spectrometer to cause charged droplets to be released from the tip of the capillary. As the droplets evaporate, gas phase charged ions are desorbed from the droplets (Aebersold and Mann 2003). ESI has advantages for constant ionization and monitoring. However, compared to MALDI, it is more sensitive to contaminating salts, buffers, and detergents. In order to fulfill ESI’s stringent requirement, high-pressure liquid chromatography (HPLC) is coupled to ESI mass spectrometry constructing a liquid-chromatography ESI-MS system (LC-MS).
One of the unique advantages of proteomics studies is the ability to analyze the post-translational modifications of proteins. Phosphorylation, glycosylation and sulfation as well as many other modifications are extremely important for protein function.
These kinds of post-translational modifications can even determine a protein’s activity, stability, localization and lifetime. The modification information is usually not coded by the gene sequence or mRNA expression data. Thereby, proteomics is the best choice to identify post-translational modification. However, this task is much more difficult than the simple protein identification which only requires minimal data such as one or two peptides to identify the protein in sequence databases. In post-translational modification analysis, all the peptides that do not have the expected molecular mass need to be analyzed. To fulfill the quantity demand, much more material is needed to study post-translational modifications than is required for protein identification.

Table of contents :

1. Intracellular transport pathways
1.1 Biosynthetic/secretory pathway
1.2 Endocytic pathways
1.2.1 Late endocytic/degradation pathway
1.2.2 Recycling pathway
1.2.3 Retrograde transport
2. Retrograde transport
2.1 The functions of retrograde transport
2.2 Pathology
2.2.1 Shiga Toxin
2.2.2 Ricin
2.2.3 Cholera toxin
2.2.4 Virus
2.2.5 Other diseases
2.3 Retrograde transport machinery
2.3.1 Clathrin coats
2.3.2 Retromer
2.4 Tools to study retrograde transport
3. Proteomics
3.1 The functions and methods of proteomics
3.1.1 Identification and analysis of proteins
3.1.2 Differential-display proteomics
a). The two-dimensional gel approach
b). Quantitative comparison by mass spectrometry
c). Protein chips
3.1.3 Protein–protein interactions
a). Purification of protein complexes
b). Yeast two-hybrid screening
c). Phage display
3.2 Proteomics in cell biology
3.2.1. A mitochondrial protein complex that links apoptosis and glycolysis
3.2.2. New component in clathrin coated vesicles
3.2.3. Kinase pathways that regulate sex-specific functions in Plasmodium
4. Reaction schemes for proteomics approaches
4.1 SNAP-tag/AGT, a DNA repair enzyme
4.2 Sulfation
4.3 Rapamycin, FK506 and FKBP
4.4 Biotin and streptavidin
4.5 N-hydroxysuccinimide in protein coupling


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