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How well can proteins discriminate between ATP and GTP? Can ATP act as a GTP mimetic?

How well can proteins discriminate between ATP and GTP? Can ATP act as a GTP mimetic?


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GTP and ATP are similar structures with the adenosine and guanosine groups differing. Both are involved in a vast array of biological functions.

However it has been shown that in certain cases, GTP and water can mimic ATP (Niefind et al., 1999).

Are there examples where ATP has acted as a mimetic of GTP? Or in other words, are there any proteins that can't discriminate between GTP and ATP very well?


Both sides now: multiple interactions of ATP with pannexin-1 hemichannels. Focus on 𠇊 permeant regulating its permeation pore: inhibition of pannexin 1 channels by ATP”

how atp and other nucleotides are released from intact cells is a fundamental question, given the existence of multiple purinergic receptor signaling cascades operative in most vertebrate tissues (25). It is well-established that neurons and neuroendocrine cells release ATP via classical mechanisms involving Ca 2+ -dependent exocytotic release of nucleotides copackaged with other neurotransmitters within specialized secretory vesicles or granules. However, many, indeed most, nonexcitable cell types locally release ATP via nonlytic mechanisms that do not involve obvious or readily measured exocytosis of nucleotide-containing vesicles or granules. An alternative mechanism for nonlytic ATP release is its facilitated efflux from the cytosolic compartment through plasma membrane transport proteins. Various membrane transport proteins or functionally characterized permeability pathways have been suggested as “ATP channels,” including some ATP-binding cassette (ABC)-family transporters, volume-regulated anion channels, plasma membrane variants of the mitochondrial voltage-dependent anion channel (VDAC) porins, and maxianion channels (31). Additionally, a strong and growing body of data indicates that ATP release from many cell types is mediated by so-called hemichannels composed of protein subunits from the well-characterized connexin (Cx) family or the recently described pannexin (Panx) family (19, 23, 26). Hemichannels can act as low-resistance conduits for the efflux of ATP and other cytosolic metabolites (59). The ubiquitous expression of the Panx1 gene in most tissues and cell types suggests that Panx1 hemichannels may comprise one of the most widely used efflux pathways for ATP release in different paracrine and autocrine signaling responses (4, 30). Notably, extracellular ATP itself, acting via certain P2Y or P2X receptors, can elicit intracellular signals that favor the gating of hemichannels to the open state. This facilitates a pathway of what can be termed “ATP-induced ATP release” linked to paracrine signaling “waves” that allow multiple cells within a tissue to respond proactively to environmental stresses (e.g., metabolic inhibition, mechanical shear, and microbial invasion) sensed by only a few cells at the immediate locus of environmental insult or stimulation (51). This sort of paracrine signaling can play a positive role in adaptive responses, such as ischemic preconditioning, relief of mechanical stress, or killing of invading pathogens, with clear physiological benefit to the whole animal. However, a cascade of P2 receptor activation coupled to the gating of ATP-permeable hemichannels also comprises a positive feedback loop that, if unrestrained, could lead to maladaptive and malignant depletion of intracellular ATP stores, collapse of ionic gradients, and cell death. Thus considerable attention has been directed toward the identification of endogenous factors that can inhibit or restrict the gating/conductance of hemichannels. In a paper by Qui and Dahl (40a), they describe a highly novel mechanism based on a direct inhibitory action of extracellular ATP on the gating and/or activity of pannexin-1 hemichannels.


Introduction

DNA molecules are the substrates for a variety of processes, including replication, transcription and recombination. Many positive regulatory mechanisms have been described which favor the selection of certain DNAs as substrates for one of these processes. In some situations, however, the positive features of a DNA can be overridden by negative regulatory mechanisms. For example, some transcriptional units are ‘silenced’ by being packaged within specialized chromatin structures which prevent interactions with RNA polymerase ( Loo and Rine, 1995 ). Eukaryotic origins of replication also become ‘silent’ after initiating DNA replication, as part of the cell cycle control that permits an origin to fire once and only once during S phase ( Muzi-Falconi et al., 1996 Wuarin and Nurse, 1996 ).

Negative regulatory mechanisms can also influence the selection of target sites in transposition. The bacterial transposons Tn3 and Tn7 and the bacteriophage Mu are sensitive to a process called target immunity, in which a target molecule that already contains a copy of one of these elements is prevented from receiving further insertions of that element ( Robinson et al., 1977 Hauer and Shapiro, 1984 Reyes et al., 1987 ). Thus, the target DNA becomes ‘silent’ or ‘immune’ to the transposition machinery.

The signal that confers immunity to a target DNA is provided by the ends of the resident Tn3, Tn7 or Mu element. Transposon ends contain special sequences that are the substrates for the DNA breakage and joining reactions that move the element from one DNA molecule to another transposon ends also contain binding sites for transposase, the enzyme which executes the DNA breakage and joining reactions (reviewed by Mizuuchi, 1992 ). The presence of Tn3, Tn7 or Mu ends in a target plasmid reduces the frequency of transposition into that plasmid 100- to 1000-fold in vivo however, transposition into other target molecules which do not contain transposon ends is not inhibited ( Lee et al., 1983 Darzins et al., 1988 Arciszewska et al., 1989 ). Therefore, target immunity is essentially a cis-acting phenomenon that prevents new insertions from occurring ‘close’ to transposon ends.

How close is ‘close’? In the case of Tn7, large (60 kb) derivatives of the Escherichia coli F plasmid are protected from Tn7 transposition when the plasmid contains Tn7 end sequences ( Arciszewska et al., 1989 ). Transposition is inhibited over even larger distances in the E.coli chromosome: the presence of Tn7 ends was shown to reduce insertions into chromosomal sites 190 kb away ( DeBoy and Craig, 1996 ). However, transposition into a target site 1.9 Mb from the Tn7 ends was not affected, demonstrating that the Tn7 ends do not cause a global inhibition of transposition ( DeBoy and Craig, 1996 ).

The ability to discriminate between targets that are ‘close’ and ‘far’ from Tn7 ends may be useful in promoting the spread and survival of Tn7. Short-range transposition events would be discouraged instead, the spread of the transposon to distant sites in the chromosome and new plasmids would be favored. Immunity would also discourage events that could potentially destroy Tn7, such as intramolecular transposition events or the hopping of one copy of Tn7 into another. Thus, target immunity plays a key role in determining what target sites the Tn7 transposition machinery will select.

The Tn7 transposition machinery also evaluates a potential target DNA for positive features. Tn7 transposition occurs at high frequency into a single site in the E.coli chromosome called attTn7 ( Barth et al., 1976 Lichtenstein and Brenner, 1982 ). Plasmids undergoing conjugation are also preferred targets for Tn7 transposition ( Wolkow et al., 1996 ). Thus, attTn7 and conjugating plasmids contain positive signals that attract the transposition machinery to these target DNAs. Different combinations of the Tn7-encoded proteins TnsA, TnsB, TnsC, TnsD and TnsE are used to select these different targets: TnsABC+D promotes transposition into attTn7, whereas TnsABC+E promotes transposition into conjugating plasmids ( Rogers et al., 1986 Waddell and Craig, 1988 ). Target immunity is observed in both the TnsABC+D and TnsABC+E transposition pathways ( Arciszewska et al., 1989 ), suggesting that the negative signal provided by a Tn7 end is dominant to the positive signals which might also be contained on a potential target molecule.

Tn7 transposition into attTn7 has been reconstituted in vitro using purified proteins ( Bainton et al., 1993 ), and the roles of the Tns proteins in executing Tn7 transposition have been investigated. TnsA and TnsB act interdependently to catalyze the chemical steps of Tn7 transposition, thus TnsA+B constitutes the Tn7 transposase ( May and Craig, 1996 Sarnovsky et al., 1996 ). TnsB binds specifically to the transposon ends ( Arciszewska and Craig, 1991 Arciszewska et al., 1991 Tang et al., 1991 ), while TnsA is likely recruited to the transposon ends through protein–protein interactions with TnsB.

The TnsA+B transposase by itself is not catalytically active TnsC, TnsD and an appropriate target DNA are also required ( Bainton et al., 1993 Gary et al., 1996 ). TnsD is an attTn7-specific DNA-binding protein which recruits TnsC, an ATPase that is also an ATP-dependent DNA-binding protein, to attTn7 targets. TnsC–TnsD–attTn7 complexes, in turn, interact with the TnsA+B transposase and activate its breakage and joining activities ( Gamas and Craig, 1992 Bainton et al., 1993 A.Stellwagen and N.L.Craig, in preparation). TnsC has been proposed to be a key connector between the target site and the TnsA+B transposase, and the ATP state of TnsC is hypothesized to regulate its ability to forge that connection ( Bainton et al., 1993 , Stellwagen and Craig, 1997 ).

Tn7 transposition occurs by a cut-and-paste mechanism, in which the element is first excised from a donor site and then inserted into a target DNA ( Bainton et al., 1991 ). The nature of the target DNA regulates both of these steps in vitro: if an attTn7 target molecule is omitted from the reaction, virtually no transposition intermediates or products are seen. Target immunity is reproduced in the in vitro Tn7 transposition reaction ( Bainton et al., 1991 , 1993 ) the evaluation of Tn7 end-containing targets also occurs early in the course of the reaction. No transposition products or intermediates are observed when the target DNA contains attTn7 but also carries a Tn7 right end. Therefore, Tn7 end-containing target DNAs are immune to Tn7 transposition not because they fail to capture excised transposons, but because they fail to provoke the excision of the transposon in the first place.

In vitro approaches have been previously used to investigate target immunity in Mu transposition. Adzuma and Mizuuchi (1988 , 1989 ) demonstrated that Mu target immunity results from the redistribution of the regulatory protein MuB from target DNAs containing Mu ends to target DNAs without ends. This redistribution is promoted by the MuA transposase, and requires ATP hydrolysis. However, it has been unclear whether this mechanism would be unique to Mu or whether it would apply to other transposons. In particular, it has been unclear how Tn7—with its multiple proteins and multiple target selection pathways—might adapt this immunity mechanism.

In this work, we have investigated the mechanism of Tn7 target immunity. We find that the key proteins responsible for Tn7 target immunity are TnsB, the transposon end-binding protein, and TnsC, the ATP-dependent target DNA-binding protein. When TnsB and TnsC are in high local concentration (i.e. when both are localized to a Tn7 end-containing target DNA), immunity is imposed on that target. An attractive model that emerges from this work is that TnsB promotes the dissociation of TnsC from Tn7 end-containing target DNAs, through an ATP-dependent mechanism. Thus, TnsB appears to impose target immunity by influencing the distribution of TnsC among potential target DNAs. We discuss the similarities between the mechanisms by which Tn7 and Mu achieve target immunity, and we discuss how Tn7 exploits this mechanism not only to avoid immune targets but also to select preferred targets for Tn7 transposition.


Cell Structure And Function QUIZ

Two types:
1) Rough ER
- "Rough" because it is studded with ribosomes --> engage in co-translational translocation
- Expanded in cells that secrete a lot (e.g. pancreatic)
- Initial site of N-linked glycosylation

Cystic Fibrosis Transmembrane Conductance Regulator (CFTR)--> Multipass ABC transporter chloride channel

Wild type --> 70% of CFTR gene produce --> misfolded --> degraded

Misfolded protein --> exposed hydrophobic regions

2) Sorting (Glyco-code)
- Glycosylated state indicates progression through endomembrane system --> used as a sorting signal for movement through the system

3) Proteolytic protection
- E.g. Mucus coat (proteoglycan)

1) Timer
- Proteins that spend too much time in ER --> likely misfolded

As protein sits in the ER --> Glucose and mannose trimming of the glycosylation occurs --> If a "core" mannose is trimmed, mannosidase (MNS1) will transport protein out of ER --> protein degradation

2) Marks for chaperone-assisted folding

Unfolded proteins are glucosylated --> glucosylated proteins are recognized by chaperone (calnexin) --> attempt fold -->>>

If folding is correct --> Protein leaves ER

COPII coat promotes movement from ER --> Golgi

Membrane proteins (like cargo receptors) have KKXX motif that binds COPI coat directly --> Budding, transport to ER

1) It can't simultaneously bind cargo and be incorporated into COPII vesicles

Highly compartmentalized --> each cisterna has a specific function (don't memorize)

1) Cis --> faces the ER --> sort traffic back to ER
2) Trans --> faces away from the ER --> transport elsewhere

1) Processing
- N-linked glycoproteins (glycosylated in the ER) undergo a series of modification to the N-linked core --> yield diverse set of sugar moieties --> multiple functions

2) Secretory vesicle
- Requires a signal --> forms a secretory vesicle --> inducable secretion (e.g. synaptic vesicles)

Lysosome microenvironment (low pH) created by V-type ATPase --> low pH activates trans-Golgi network derived acid hydrolases (inactive in neutral pH)

2) Autophagosomes
- Double membrane-bounded organelles formed around defunct organelles or in nutrient-starved cells

M6P receptor is retrieved -->>>>

1) Constitutive
- Does not require specific signal --> Always occurring (cargo or not)
- Replaces membrane components lost by endocytic processes

1) Phagocytosis --> materials greater than 250 uM in size --> clathrin-based

Cholesterol is packaged as LDL particles when carried in the blood

2) Transcytosis (requires signal)
- Enter the recycling endosome --> Transferred to the other side of the cell

These are essentially vesicles that form inside endosomes --> quarantine transmembrane catalytic activity of receptors as well as allow access to cytosolic catalytic regions that would otherwise not be in the lumen

2) Surface membrane retrieval (Endocytic pathway)
- Involves uptake of macromolecules --> for biosynthesis and recycling membrane components

Due to the large amount of vesicles --> Less dense

E.g sex1 cells accumulate vesicles, sec 22 have engorged ER, what do sec1/sec22 double mutants look like

If double mutant looks like sec1 --> accumulation of vesicles occurs first

If double mutant looks like sec22 --> engorged ER occurs first

After fission --> Hydrolysis of GTP --> Molecules dissociate

Two parts used in Combinatorial Recognition:

1) BAR domains - are sensitive to membrane curvature --> higher affinity for highly curved regions of the membrane

Binding of v-SNARE with t-SNARE dehydrates polar head groups --> excludes water

SNAREs linked to the membrane with lipid anchors --> fusion fails to go to completion

Transmembrane domains of SNARE are important --> protein shape can change lipid composition and packing and overall membrane curvature

Oleic Acid --> cone-shaped --> stimulate membrane fusion

1) Wavelength of the light

Low illumination --> small amount of particles --> obscured by statistical fluctuations

TIssues need to be:
1) fixed --> cross links all proteins (formaldehyde) --> permeates cell (opens holes to allow dye in)
2) Dehydrated --> increases shelf life (alcohol)

Hematoxylin (Blue) --> binds arginine/lysine --> Nuclei

size of cell organelles <0.5 um --> need something strong

Two types:
1) Charged couple device cameras (CCDs)
2) Complmentary metal-oxide semiconductor sensors (CMOS)

In microscopy, a fluorescent marker is attached to the antibody --> allowing visualization/localization of that target antigen

Visualization of internal structure requires fixation to permeabilize the membrane --> allows antibody into the cell

Used as a reporter molecule
--> gene can be attached to another gene (fusion protein)
--> protein can be attached to an antibody

Uses computer algorithms to reduce the optical diffraction of light coming in from both the plane of focus as well as the planes above and below the focus (although still constrained by the diffraction limit)

Another pinhole aperture is placed in front of the detector (confocal with the illuminating pinhole) --> precisely where the rays emitted from the specimen is focused

Preactivation- absorption peak in the 400 nm range --> absorption of this wavelength activates it

Uses Photoactivated (PA)-GFP

Uses lasers to sequentially switch on (ex: near-ultraviolet light) a sparse subset of fluorescent molecules --> Images --> Bleaches subset --> activates next subset

Works in vacuum
Uses magnets as "lens"

Proposes that at one point in history, one prokaryote engulfed (phagocytosis) another prokaryote

Proposes that the invagination of the outer membrane --> Nucleus

One or more nucleoli per eukaryotic nucleus --> represents high levels of rRNA synthesis required for cellular growth

Results in anemia from two possible mechanisms:
1) Activation of tumor suppressor gene TP53 --> apoptosis of erythroid progenitors --> anemia

Function in structural integrity of the nucleus, organization of pores and gene regulation (via pore regulation

Highly compacted and rich in histone modifications associated with repression

The most active DNA sequences are gated near the nuclear pore

1) GFP-labeled Lac repressor with a FKBP3 protein attach
2) eGFP-lac repressor-FKBP3 protein binds to --> Operator of the Lac operon --> Operon repressed --> no gene expression
3) Addition of rapamycin allows binding of FKBP3 --> to the FRB domain of mTOR (which contains transcriptional activator VP16)
4) Heterodimerization of FKBP3 to FRB --> allows transcription factor VP16 to exert activity on the Operon --> Operon is expressed --> gene expression is ON

No Rapamycin --> no gene expression --> GFP labeled loci remains in the peripheral

These signals are often found at the N-terminus of the polypeptide chain --> many of which are removed by signal peptidases once sorting process is complete

Each one contains aqueous passages --> allows small water-soluble molecules to diffuse

Rate of diffusion is governed by size of these molecules (smaller are quicker)

All share a common characteristic --> have FG (Phenylalanin-Glycine) repeat proteins --> Hydrophobic in nature


Significance of Ras Signaling in Cancer and Strategies for its Control

Ras is a GTP-binding protein and is the most widely studied oncoprotein. To achieve its biological activity, it must undergo post-translation modification. Ras acts as a typical molecular switch. The GTP-bound Ras can activate several downstream effector pathways. Ras signaling regulates many important physiologic processes within a cell, such as cell cycle progression, survival, apoptosis, etc. Several studies have found mutation in Ras or its effectors in various types of tumors. Therefore, Ras or its downstream effectors can be attractive drug targets against various types of tumors in cancer therapeutics. Some therapeutic agents against Ras effectors, such as Raf, MEK1/2, PI3K, AKT etc., have successfully managed to enter into phase I and II trials. This targeted drug design could be envisaged in mainly four ways, such as prevention of Ras-GTP formation, covalent locking of the GDP-bound Ras, inhibition of Ras-effector interactions, or impairment of post-translational modification of Ras. In this review we summarize the normal Ras signaling as well its aberrant signaling in tumors and various strategies to inhibit Ras signaling.

Keywords

Ras signaling, oncoprotein, effectors, apoptosis

Article:

H-Ras, K-Ras, and N-Ras are the main members of Ras superfamily, which bind small molecules GTP and GDP interchangeably and can hydrolyze GTP to GDP. The Ras superfamily consists of more than 150 proteins and these can be classified under at least five sub-families viz: the Ras, Rab, Rho, Ran, and Arf families. 1 Ras oncoproteins act as typical molecular switch by alternately binding to GTP and GDP molecule and has intrinsic GTPase activity. It remains in an active state when bound to GTP and switches to an inactive state by binding to GDP and thus controls the expression of the downstream genes. Ras signaling is an important intracellular signaling pathway that plays a role in cellular proliferation and differentiation, survival, and gene expression. 2–4 Ras oncoprotein has also been implicated in the development of cancer by either having increased intensity or prolonged signaling mechanism. 5 This may happen either due to a mutation in the Ras-GTPase domain, which renders it constitutively inactive (GDP-bound state), or due to activating mutation in growth factor receptors that act upstream of Ras or due to aberrant RAS effector activation. Due to the elevated level of Ras signaling in tumor growth and progression, Ras proteins and its downstream effector proteins may serve as promising therapeutic targets against cancer. In this review article we shed light on normal Ras signaling as well as its aberrant signaling in tumors and proposed various strategies for its inhibition.

Normal Ras Signaling Pathways
The three human RAS genes encode four proteins with a size of

21 kDa: H-Ras, N-Ras and the splice variants K-Ras4A and K-Ras4B. A newly synthesized Ras protein is a soluble cytoplasmic protein, which needs to undergo post-translation modifications to associate with particular lipid membrane. These modifications occur at the carboxyl terminal ‘CAAX’ box (denoting amino acid sequence cysteine-aliphatic-aliphatic-X residue). The first step involved is the attachment of 15 carbon farnesyl to the cysteine residue of the ‘CAAX’ box.6 Next, an endopeptidase Rce1 cleaves off three terminal amino acid residues and the resulting isoprenylated cysteine residue is methylated by the isoprenylcysteine carboxyl methyltransferase (ICMT) as shown in Figure 1. 7 H-Ras and K-Ras undergo additional palmitoylation modification.8 These post-translational modifications are necessary for binding to lipid membrane to execute their biologic functions.

The three-dimensional structures of Ras proteins with bound GTP and GDP and their mutant variants were determined through X-ray crystallography in 1990. 9–11 The Ras protein consists of a hydrophobic core of six β sheets and five α helices that are interconnected by a series of 10 loops. Five of these loops determine the high-affinity nucleotide interactions of Ras and regulates GTPase activity. The GTP γphosphate is stabilized by interactions that are established with the residues Lys16, Tyr32, Thr35, Gly60, and


Gln61 of loops. Gln61 is a key residue that stabilizes the transition state of GTP hydrolysis to GDP, in addition to participating in the orientation of the nucleophilic attack that is necessary for this reaction. It has been seen that oncogenic mutations of Gln61 reduce the intrinsic GTP hydrolysis rate, thereby rendering the Ras protein constitutively active. The structural differences in GDP-bound Ras (inactive state) and GTP-bound Ras (active state) lies mainly in highly dynamic regions termed Switch I (residues 30–40) and Switch II (residues 60–70) as shown in Figure 2, which are required for the interaction of Ras with both upstream as well as downstream partners. The binding of GTP brings change in structural conformation of side chain of Switch I, via inward reorientation of side chain of Thr35, which facilitates its interaction with the GTP-γ phosphate as well as the Mg2+ ion. Similarly, the γ-phosphate induces significant changes in the conformation of Switch II through the interaction it establishes with Gly60. 12

Besides its normal function, such as cellular proliferation and differentiation, survival, and gene expression, gain-of function mutations of H-, N-, K-Ras have been found in different types of human cancers. 12,13 Besides, aberrant Ras signaling is also implicated in several developmental disorders, known as the cardio-facio-cutaneous diseases (i.e., neurofibromatosis-type I [NF-1], Costello syndrome, and Noonan syndrome). 14

The normal biological functioning of RAS proteins as described before require post-translational modification, guided by many important enzymes described earlier. These RAS preprocessing enzymes can serve as attractive drug targets. The activity of RAS is largely determined by the type of cofactors that it binds. When it is bound to GTP, it is active and can recruit downstream target proteins, but when it binds GDP, it is rendered inactive and fails to interact with the downstream effectors. This association of RAS with GTP or GDP is mediated by two enzymes: guanine nucleotide exchange factors (GEFs) and GTPase-activating proteins (GAPs). GEFSs catalyze the exchange of GDP for GTP whereas GAPs increase the rate of GTP hydrolysis to GDP plus phosphate. 15

The activated RAS protein can bind and activate downstream effectors, which ultimately produce appropriate signals, such as cell proliferation, survival, and other physiological functions through the activation of various pathways (see Figure 3).

One of the first mammalian effector of RAS, which has been well studied and characterized is the protein serine/threonine kinase, is RAF. 16,17 There are three closely related RAF proteins, CRAF1, BRAF, and ARAF, which are known to be activated by RAS-bound GTP.18 The activated RAF can phosphorylate and activate the downstream targets, such as mitogenactivated protein kinases 1 and 2 (MEK1 and MEK2).19 MEK1 and MEK2 can then phosphorylate and activate mitogen-activated protein kinases (MAPKs), such as extracellular signal-regulated kinases 1 and 2 (ERK1 and ERK2). The substrates for ERK1/2 include both nuclear as well as cytosolic proteins, of which transcription factors have been widely studied. 20,21 ERK phosphorylate (ELK1) belongs to ETS family transcription factors, which, in turn, regulates the expression of FOS. 22 Additionally, ERK can also phosphorylate c-JUN. 23 The activation of all these transcription factors eventually promotes the cell cycle progression. 24

In addition to the RAF/MAPK effector pathway, RAS can also interact with another well-characterized effectors ie., phosphatidyl inositol 3-kinases (PI3Ks). 25,26 The activated PI3Ks phosphorylate phosphatidylinositol 4,5-bisphosphosphate (PtdIns(4,5)P2) produce a second messenger phosphatidylinositol- 3,4,5 -triphosphate(PtdIns(3,4,5)P3), which interacts with several other proteins through pleckstrin homology and other domains. 27 Thus, PI3K exercises its control on a large number of downstream target proteins. PI3K regulates the activity of two important kinases PDK1 (3-phosphoinositide-dependent protein kinase-1) and AKT. 28,29 PDK1 is important for the activation of many protein kinases of the AGC family, comprising AKT/PKB, p70S6K, some PKCs and RSKs. 30–33 AKT has an antiapoptotic function and have been found to be important for survival signals generated by RAS. 34,35 Besides, PI3K activation also leads to the activation of RAC, a RHO family protein that not only regulates the actin cytoskeleton but also transcription factors, such as nuclear factor-kappa B( NF-κB). 36–38 RAC activation also seems to be important in RAS-induced transformation. 39,40

The third well-known effector of RAS include three exchange factors for RAS-related RAL proteins: RAL guanine nucleotide dissociation stimulator (RALGDS), RGL2/RLF, RALGDS-like gene (RGL/RSB2). 41–43 With the help of these proteins RAS is able to activate RAL, which in turn activates phospholipase D1, CDC42/RAC-GAP-RAL binding protein1 (RALBP1). 44,45 These RALGDS pathway together with AKT contribute to the inhibition of FORKHEAD transcription factors. 46 These have been involved in arresting cell cycle progression through activation of cyclin dependent kinase inhibitor, KIPI (also known as p27), and apoptosis through expression of the BIM and FAS ligands. 47,48

Phospholipase Cε is another effector of Ras. 49 It hydrolysis phosphatidylinositol 4,5-bisphosphosphate to diacylglycerol and inositol-1,4,5-triphosphate, which leads to activation of PKC and calcium mobilization. 50

Through the concerted efforts of RAS and its effectors, it is able to regulate a wide array of functions, such as cellular proliferation, survival, apoptosis, and other important physiological processes.

Abnormal RAS Signaling in Tumors
Activating Mutation in RAS

The aberrant RAS signaling in tumors can be contributed by several different mutations, mostly activating mutation in tumor cells: 85 % in K-RAS, 15 % in N-RAS, and 10 %) was found in tumors, such as colon (adenocarcinoma), leukemias (AML), lymphomas (Hodgkin’s lymphoma), lung (large-cell carcinoma and non-small-cell carcinoma) and thyroid (anaplastic and folllicular carcinoma). It is also evident from Table 1 that the oncogenic mutation predominately affects the K-Ras locus, with oncogenic K-Ras mutations being detected ranging from 16 to 36 % in various human tumors screened. The activating mutations mostly affect the GTPase activity of RAS leading to accumulation of RAS-bound GTP. 51,52 These GTP-bound RAS can hyperactivate other downstream effector proteins previously discussed before leading to constitutive abnormal signaling and anarchy within the tumor cell. The impaired ability of Ras mutants to hydrolyze GTP, either intrinsically or in response to GAPs, is responsible for the oncogenic nature of mutations at residues G12, G13, and Q61 in the active site. 53

Ras remains activated due to loss of GAP-accelerated GTP hydrolysis. One such typical example of GAP mutation is the GAPs, neurofibromin encoded by the NF1 tumor suppressor gene. 54 Patients with neurofibromatosis type I inherit onlyone functional NF1 gene and then predisposed to cancer through complete loss of NF1.

Growth Factor Receptor Activation
Ras signaling has also been known to be activated in tumors in which growth factor receptor tyrosine kinase has been overexpressed. The most common example are epidermal growth factor receptor (EGFR) and receptor tyrosine-protein kinase erbB-2 (ERBB2) which are activated and overexpressed in many types of cancer including breast, ovarian, and stomach carcinomas. 55

Mutation or Amplification of Ras Effectors It has been found that BRAF is commonly activated by mutation in human tumors, such as melanomas and colon carcinoma. A study of 923 cancer samples reveal that missense mutations occur commonly in the BRAF


gene in approximately 70 % of human malignant melanoma and 15 % of colorectal cancers. 56 The major mutation found in tumors is V559E, a substitution of valine by glutamic acid, which occurs in the kinase activation domain resulting in activation of BRAF. 56

The PI3K pathway is known to be activated due to amplification or mutation of PIK3CA encoding catalytic p110α subunit of PI3K in ovarian and cervical tumors. 57,58 These mutations commonly occur in two conserved regions of the gene, which encode the kinase and helical

domains of the protein. These ‘hot spot’ mutations, H1047R, E545K, and E542K, are nonsynonymous missense mutations that confer constitutive kinase activity. 59,60 Second mechanism of PI3K activation occurs by amplication of its downstream target AKT2 in ovarian and breast tumors. 61 A somatic missense mutation in the pleckstrin homology (PH) domain of AKT1 (E17K) had been identified in breast, colorectal, and ovarian cancers. 62 Additionally, PI3K can be directly activated due to loss of tumor suppressor gene PTEN (phosphatase and tensin homolog deleted on chromosome ten). This gene encodes a lipid phospholipase that dephosphorylate phosphatidyl-3,4,5-triphosphate at position 3 of the inositol ring, which reverses the accumulation of these second messengers caused by PI3K and thus negatively regulates PI3K activity. PTEN is one of the most frequently mutated genes in human cancers. 63,64

Ras Inhibition Strategies

The involvement of aberrant Ras signaling in different kinds of tumors in humans necessitates the development of therapeutic agents, which can restore a normal function in tumor cell. The developments of therapeutics against it have already started and some have successfully managed to enter into phase I–II clinical trials (see Table 2). Targetting the aberrant Ras signaling has largely come from indirect inhibition of Ras effectors, such as AKT, MEK-1/2, PI3K, Raf, etc. This Ras signaling inhibition strategies can be categorized into following four inhibition strategies (Figure 4) each one is discussed briefly below.

Prevention of Ras-GTP Formation
Early studies explored the potency of ATP-competitive kinase inhibitors, which were expected to be used for inhibiting nucleotide binding to Ras. Although the affinity of kinases for ATP are usually in the micromolar range, 65 picomolar nucleotide affinities of Ras combined with millimolar intracellular nucleotide pools complicate the use of GTP-competitive inhibitors. 66 Consequently, strategies that prevent the initial formation of the Ras–GTP complex were investigated.

Libraries of GTP analogs with alternations at the ribose or nucleotide moiety 66 and of pyrazolo[3,4-b]quinoline ribosides 67 yielded molecules with moderately increased affinity compared with GDP and relatively weak inhibitory potency.

Covalent Locking of the GDP-bound State

Because of the high affinity of Ras to GDP, nucleotide exchange requires assistance by GEF proteins such as Son of Sevenless (SOS) or the closely related RasGRF1 to facilitate GTP loading. Early examples of GEF inhibitors were identified from a compound library whose members were originally designed to compete with GDP for nucleotide binding. SCH53239 and its derivatives target a hydrophobic pocket close to the nucleotide binding site and inhibit the intrinsic nucleotide exchange. 68,69 On the basis of these studies, a series of derivatives with improved potency and water solubility was designed that is capable of inhibiting the RasGRF1-catalyzed nucleotide exchange in vitro (half-maximum in hibitory concentration [IC50] = 35−320 μM). 70,71 The orthosteric Ras-GEF interaction inhibitors described above do not discriminate between mutated and wild-type Ras. In an approach to overcome this limitation for mutated K-RasG12C, the thiol function of cysteine 12 was used to covalently trap inhibitors. A set of GDP-derived inhibitors was developed to directly target the nucleotide binding site. 72 The most active compound, SML-8-73-1, covalently binds K-RasG12C even in the presence of millimolar concentrations of GDP and GTP. SML-10- 70-1 shows antiproliferative activity in Ras-dependent cells expressing K-RasG12C (EC50 = 27−47 μM). 73

Ostrem et al. 74 reported covalent inhibitors that are selective for mutant cysteine over the wild type as it relies on trapping of the thiol group in Cys12 in common oncogenic mutant (K-Ras G12C) using disulphide-l fragment-based screening approach. Binding of these inhibitors to K-Ras (G12C) destabilize the native nucleotide preference to favor GDP over GTP and impedes binding to Raf.

Inhibition of Ras-effector Interactions
Protein–protein interfaces (PPIs) between Ras-GTP and effectors initiate various downstream signaling cascades. Remarkably, Ras–GTP exists in at least two distinct conformational states, which interconvert with rate constants on the millisecond timescale. 75 Whereas state 2 represents a conformation with high affinity for effector binding, the affinities for effectors exhibited in state 1 are reduced. 76,77 However, state 1 exhibits surface cavities potentially accessible to small molecules that could stabilize this conformation and thereby inhibit interactions with effector proteins. Zn2+-cyclen, Cu2+-cyclen, and bis(2-picolyl)amine complexes bind H-Ras–‘GTP’ with millimolar affinity and stabilize the ‘noneffector binding’ state 1. 78,79 The feasibility of orthosteric Ras-effector interaction inhibition was shown with antibody fragments that block effector interaction sites of H-Ras–GTP. 80 In this setup, disruption of mutant Ras-effector interactions is sufficient to prevent tumor initiation in a transgenic mouse model of lung cancer. 81 Small peptide–based inhibitors lacking drug-like properties have been used to disrupt Raseffector interactions. 82,83

Impairment of Post-translational Modification of Ras
Ras-dependent signaling requires the correct intracellular localization of Ras proteins predominantly at the plasma membrane, mediated by membrane-anchoring lipid residues at the C terminus. Therefore, impairment of Ras localization has been explored to inhibit oncogenic Ras signaling. 84 Ras proteins are equipped with lipid groups through a series of post-translational modifications, which include cysteine S-farnesylation, proteolysis, and carboxymethylation at the C terminus of all Ras isoforms and additional cysteine S-palmitoylation of H-Ras and N-Ras. 85

FarnesyltransfeRase inhibitors (FTIs) interrupt this biosynthetic sequence, leading to nonlipidated cytosolic Ras. Several FTIs reached late-stage clinical trials but ultimately failed, mostly because of alternative geranylgeranylation of the K-Ras and N-Ras isoforms. 86 Treatment with

geranylgeranyl transfeRase inhibitors 87 or dual prenylation inhibitors 88 did not show clinical efficacy. However, a number of studies report promising preclinical and clinical results for FTIs as single agents or in combination with other conventional anticancer agents. 89 FTIs may also show positive clinical responses in the treatment of H-Ras-dependent cancers and tumors that rely on other farnesylated proteins for survival. 87

Conclusion
Ras signaling pathways constitute central drivers of cancer development and therefore strategies have been sought for development of potent Ras inhibitors effective in vivo as well. Some inhibitors against Ras effectors such as Raf, MEK1/2, PI3K, AKT, etc. have already entered into clinical trials, which suggests the relevance of Ras signaling pathways in cancer therapeutics. Since Ras signaling pathway is a complex network controlled by several feedback loops, blocking a single pathway may not be adequate to achieve a reduction of Ras signaling to a therapeutically significant level. Therefore, combined therapies targeting Ras and other oncoproteins in parallel may be required to control tumors.

Article Information:
Disclosure

Arun Bahadur Gurung, MSc, and Atanu Bhattacharjee, PhD, have nothing to disclosure in relation to this article. No funding was received in the publication of this article.


Materials and Methods

Materials

Anti-SRP54 was a gift from B. Dobberstein (ZMBH, Heidelberg, Germany), whereas anti-Hsp40 and anti-Hsp70 antibodies were from Stressgen. Canine SRP was prepared using established protocols (Walter and Blobel, 1983b), but omitting low concentrations of the detergent Nikkol in any buffers. Canine pancreatic microsomes (Walter and Blobel, 1983a) were depleted of endogenous SRP (supplementary material Fig. S1A) by washing in high-salt buffer (Walter and Blobel, 1983b). Hsc70 was purified from bovine brain by chromatography on DEAE-cellulose, ATP-agarose and hydroxyapatite (supplementary material Fig. S2). Recombinant human Hsp40 and Hsp90 were obtained from Stressgen. The C-terminal domain of human Bag-1M [C-BAG, residues 151-264) in the vector pPROEXHTa (Invitrogen)] was expressed in BL21(DE3) E. coli and purified by Ni-Sepharose and Mono Q chromatography (supplementary material Fig. S2) as previously described (Sondermann et al., 2001).

Transcription

cDNAs encoding human Sec61β and rat synaptobrevin 2 were cloned in to pSPUTK (Abell et al., 2004) and transcription templates incorporating a C-terminal glycosylation tag or replacing the hydrophobic tail-anchor region were prepared by PCR using appropriate reverse primers (see supplementary material Table S1). Sec61βOPG was created in pCDNA5 (Invitrogen) by mutagenesis and the transcription template obtained by PCR from the resulting construct (supplementary material Table S1). In all cases, the mRNAs lacked a stop codon causing the resulting polypeptides to remain associated with the ribosome after synthesis (see Fig. 1A for protein sequences). Transcripts were synthesised using SP6 or T7 RNA polymerase, according to manufacturer's instructions (New England Biolabs or Promega, respectively).

Translation and membrane insertion

Proteins were synthesised using rabbit reticulocyte lysate with incubations at 30°C in the presence of [ 35 S]-methionine, according to manufacturer's instructions (Promega). Puromycin was used at 1 mM with subsequent incubation at 30°C for 5 minutes to elicit efficient release of the stalled peptidyl-tRNAs from the ribosome (Abell et al., 2004). SRP-depleted microsomes (K-RM) were added to a final concentration of 1.5-2.0 OD280 per ml, and were analysed for TA protein insertion on the basis of relative N-glycosylation efficiency following recovery by centrifugation through 100 μl HSC (500 mM sucrose, 500 mM KOAc, 5 mM Mg(OAc)2, 50 mM Hepes-KOH pH 7.9) at 100,000 g for 10 minutes or 132,000 g for 5 minutes. Where indicated, the resulting membrane pellet was resuspended in 100 μl of cold 0.1 M Na2CO3, incubated on ice for 10 minutes and recovered by centrifugation at 132,000 g for 5 minutes to confirm membrane integration. De-glycosylation was performed with endoglycosidase H (EndoH) according to manufacturer's instructions (New England Biolabs).

Nucleotide depletion

Reticulocyte lysate was depleted of nucleotides by loading 70 μl onto a Biospin 6 column (Bio-Rad) equilibriated with LSC buffer (100 mM sucrose, 100 mM KOAc, 5 mM Mg(OAc)2, 50 mM Hepes-KOH pH 7.9, 1 mM DTT), following manufacturer's instructions, repeating the process once. A parallel depletion using a translation of Syb2 showed a 49% recovery rate and a double volume of depleted lysate was used for comparative experiments with non-depleted lysate.

Crosslinking and immunoprecipitation

Following puromycin treatment, translation products were treated with 1 μg of apyrase per 40 μl volume for 5 minutes at 30°C, then incubated on ice for 5 minutes followed by incubation at 30°C for 5 minutes with either 1 mM disuccinimidyl suberate (DSS Pierce), 1 mM succinimidyl trans-4-(maleimidylmethyl) cyclohexane-1-carboxylate (SMCC Pierce) or bismaleimidohexane (BMH Pierce) diluted from a 20 mM stock in DMSO. Crosslinking was stopped with 50 mM glycine (DSS), 10 mM 2-mercaptoethanol (BMH) or both (SMCC). Samples were denatured with SDS unless otherwise stated specific adducts were recovered by immunoprecipitation (Abell et al., 2003).

Reconstitution of ER integration

Ribosome–nascent-chain complexes (RNCs) were generated by translating transcripts lacking a stop codon for 7 minutes. Reactions of 200 μl were supplemented with 2.5 mM cycloheximide and 500 mM KOAc, and the final 240 μl sample was layered over 400 μl HSCC (HSC with 2.5 mM cycloheximide and 1 mM DTT), followed by centrifugation at 213,000 g for 20 minutes. The pellet was resuspended in 50 μl HSCC with reduced sucrose (100 mM), layered onto 150 μl HSCC, and centrifuged at 213,000 g for 20 minutes. The pellet was finally resuspended in 40 μl LSC. Membrane-insertion reactions comprised 2 μl of isolated RNCs made up to a final volume of 10 μl by LSC and various additions. Hsp40 was added at 3 μM, Hsc70 was added at 1.7 μM, Hsp90 was added at 1.3 μM, TRiC (gift from Judith Frydman, James Clark Center, Stanford University, CA) was added at 0.6 μM, SRP was added at ∼12.5 nM, prespun reticulocyte lysate was added at 20% v/v, and depleted lysate was added at 40% v/v. ATP or GTP was added at 1 mM. Following the addition of all cytosolic targeting factors and treatments, puromycin was added at 1 mM and the sample incubated for 5 minutes at 30°C. Membrane insertion was achieved by incubation with K-RMs (final concentration of 1.5-2.0 OD280 per ml) at 30°C.

Gel electrophoresis

Samples were heated to 70°C for 10 minutes in SDS-PAGE sample buffer and then resolved on 16% polyacrylamide Tris-glycine gels under denaturing conditions. Gels were fixed, dried and then exposed to phosphorimage plates, which were read using a Fuji BAS-3000 phosphorimager. Radiolabelled products separated by SDS-PAGE were quantified using Aida software.


Biological switches and the second law of thermodynamics

One could argue that the concept of a singular ON or OFF state in a molecular switch might violate the Second Law of Thermodynamics. The Second Law requires that biochemical systems transit one state to the other by a series of microscopically reversible steps. This idea is based in statistical mechanics as it is applied to a system at equilibrium, which must be applied a priori to enzyme catalyzed biological processes. It is easy to visualize the origins of the principle of microscopic reversibility by considering the consequences were it NOT true. For example, if the rate of A → B were greater than B → A at equilibrium, each of the rates B → C, C → D, and D → A would also have to be greater than their reverse rates to prevent buildup of the concentration of any species, which is not permitted at equilibrium. In this case there would be a preferred direction-of-operation of the reaction cycle. Such a spontaneous cycle in a system at equilibrium (i.e., an engine that spontaneously produces work) is not consistent with the drive toward maximum entropy contained in the Second Law.

There is no violation of the Second Law if the transit from an OFF to ON state (or vice versa) occurs reversibly. The molecular basis for this type of microscopic reversibility can be visualized for the hMSH2–hMSH6 and G-protein switches as reversible nucleotide binding as well as intermediate protein conformational changes that occur while transiting the extreme states. It is these conformational transitions that determine interaction with effectors which is ultimately paid for by the hydrolysis of NTP. More significantly, one can affect the equilibrium of each state experimentally by altering the ratio of NDP/NTP in the absence of any hydrolysis (see Fig. 6 inGradia et al. 1997). It is also important to note that microscopic reversibility has been directly demonstrated for the gated maxi K + ion pump, a molecular switch controlled by similar conformational transitions (Song and Magleby 1994). Thus, molecular switches are both reversible and, at equilibrium, clearly preserving a fundamental tenant of thermodynamics.


Role of RNA in Protein Synthesis | Microbiology

The processed RNA molecules take part in protein synthesis with the help of ribosomes.

All the three types of RNAs are involved in protein synthesis in the following main steps:

(i) Activation of amino acids,

(ii) Transfer of amino acid to tRNA,

(iii) Initiation of protein synthesis,

(iv) Elongation of the polypeptide chain.

During the process of translation the genetic information’s are coded in mRNA transcripts in the form of codons which in turn are specifically read by anticodon of tRNA and used to form a polypeptide molecule of defined function.

1. Charging of tRNA:

(i) Activation of Amino Acids:

In protein synthesis only L-amino acids take part. The D- amino acids are screened from the all 20 amino acids. In addition, the other amino acids which are not used in protein synthesis are citrulline, alanine, β-alanine, etc. Each amino acid has a specific aminoacyl tRNA synthetase (charging enzyme) and a specific tRNA. At least 32 tRNAs are required to recognise all the amino acid codons, but some cells used more than 32.

However, these may be more than one species of tRNA for a specific amino acid but there is only one charging enzyme for each amino acid. Its carboxyl group activates the amino acids through being catalysed by its own specific activating enzyme (aminoacyl tRNA synthetase) in the presence of ATP. Consequently aminoacyl AMP synthetase complex is formed which remains in bound form with the activating enzyme.

(ii) Transfer of Amino Acid to tRNA:

The process of transfer of activated amino acids to tRNA is called charging of tRNA. The tRNAs are specific to their specific amino acid. Therefore, tRNAs are named according to specific amino acid such as tRNA ala (for alanine), tRNA val (for valine), etc. Therefore, the activated amino acid is transferred to its specific tRNA.

The aminoacyl- AMP-synthetase complex formed as above is transferred to tRNA as below:

Aminoacyl-AMP- synthetase complex + tRNA → Aminoacyl – tRNA + AMP + aminoacyl tRNA synthetase Structure of aminoacyl tRNA is given in Fig. 10.14. The aminoacyl-AMP synthetase reacts with specific tRNA and forms aminoacyl-tRNA complex by releasing the enzyme aminoacyl- tRNA synthetase.

This shows that the enzyme tRNA synthetase has two specific sites. One site recognises the specific amino acids and the other site recognises the specific tRNA molecule. Thus, the tRNA synthetase brings the specific amino acid and tRNA molecule together.

However, these recognition properties are essential for making sure that the specific amino acid is charged on the proper tRNA molecule. In the same way the tRNA molecule also consists of two specific sites, one site for recognising its specific aminoacyl- tRNA synthetase and the second (the anticodon) for codon present on mRNA molecule.

For the incorporation of an amino acid at proper position in the polypeptide chain, recognition of codon on mRNA by the specific anticodon on tRNA is required.

2. Initiation of Polypeptide Synthesis:

There are several specific and complex processes (Fig. 10.15) that are involved in the initiation and continuation of the elongation of polypeptide sequence. The essential components required for initiation are: initiation factors, ribosome, mRNA, guanosine triphosphate (GTP) and aminoacyl-tRNase.

Fig. 10.15 : Initiation of translation.

(i) Initiation Factors:

There are certain initiation factors (IF) which are required for the initiation of protein synthesis. In prokaryotes three IF (i.e.IF-1, 9,000 MW IF-2, 1,15,000 MW and IF-3,22,000 MW) are involved in the initiation process, whereas in eukaryotes no IF equivalent to IF-1 and IF-2 are found.

However, IF-2 is functionally equivalent to eukaryotic eIF-2 and elF- 2′ and IF-3 is equivalent to eukaryotic eIF-3. The IF-1, IF-2 and IF-3 are present in the 30S subunit of the ribosome. IF-1 and IF-2 help in binding of initiation tRNA (tRNA met ) to the 30S ribosome subunit.

(ii) Formylation of Methionine:

Methionine is the starting N-terminal amino acid in eukaryotes, whereas in prokaryotes methionine consists of a formyl group (-CHO). Therefore, formyl group containing methionine is called N-formyl methionine. In prokaryotes as well as in eukaryotes initiation of protein synthesis occurs through a specific methionyl tRNA which is commonly known as initiation tRNA (i.e. tRNA met ).

Binding of initiation tRNA with methionine/formylmethionine occurs as below:

Methionine + tRNA → Methionine – tRNA (met – tRNA)

Formyl tetrahydrofolate + NH2-methionyl tRNA Transformylase → N-formyl-methionyl-tRNA (N-fmet-tRNA)

(iii) Formation of 30S Initiation Complex:

The first step in initiation of protein synthesis is the formation of 30S initiation complex. This complex consists of an mRNA, 30S ribosomal subunit, GTP, IF (1, 2 and 3) and the initiator tRNA i.e. N-fmet-RNA.

Formation of 30S initiator occurs in the following steps (the actual order of these steps is not known):

(a) The initiation factors (IF-1, IF-2 and IF-3) bind to 30S ribosomal subunit in the presence of GTP to form 30S-IF complex (Fig. 10.15A). However, when the mRNA is absent IF-1 and IF- 3 do not form complex neither with 30S subunit nor 50S subunit.

(b) The second step involves the association of mRNA and initiator tRNA to the 30S subunit. However, the actual order of these steps vary. The IF-3 can bind to both 30S subunit and to mRNA. The 30S-IF complex binds to mRNA at the site containing initiation codon (in the order of pB reference AUG, GUG, UUG, CUG, AUA or AUU). Each mRNA at its un-translational region consists of a ribosome binding site for every polypeptide in the form of polycistronic message.

This ribosome binding site (i.e. 5′-AGGAGGU-3′) is known as Shine-Dalgarno sequence which is important in the binding of mRNA to the 30S-IF complex (Fig. 10.15B). The Shine-Dalgarno sequence base pairs to a region present at 3′ end of 16S rRNA. This pairing will results in proper position of initiating AUG codon so that it can combine with an initiator anticodon on tRNA.

(c) The IF-2 which has combined with GTP, permits the initiator tRNA (i.e.N-fmet-tRNA) to bind to the 30S ribosomal subunit (Fig. 10.15C). Then it allows to the 30S and 508 subunits to get associated. This binding is followed by removal of IF-3 from the 30S-IF complex.

Removal of IF-3 is necessary because its presence inhibits the association of two ribosomal subunits. At this stage the initiation complex consists of mRNA associated with the 30S ribosomal subunits, IF-1, IF-2-GTP and fmet-tRNA.

(iv) Formation of the Complete Initiation Complex:

The last step in prokaryotes is the union of the 30S initiation complex with 50S ribosomal subunits and formation of a complete 708 initiation complex (Fig. 10.15D). This process of union causes the immediate hydrolysis of the bound GTP to GDP + Pi.

The process of union is accomplished in the presence of an analogue of GTP (i.e. 5’guanyl methylenediphosphate). Therefore, hydrolysis of GTP and subsequent removal of GDP is essential for the IF-1 dependent release of IF-2 from the ribosome (Fig. 10.15 E). Similarly, in eukaryotes the 40S initiation complex is attached to 60S ribosomal subunit and forms the complete 80S initiation complex.

The ribosome has three important binding sites, two are important in protein synthesis. The two binding sites are: the aminoacyl-tRNA binding site (A), the peptide (P) binding site and the E site (Fig. 10.15E).

The A site receives all the incoming charged tRNA, whereas the P site possesses the previous tRNA with the new polypeptide (peptidyl tRNA) attached. The fmet-tRNA (initiation tRNA) directly binds with P site, but not A site. Function of the E site is de-acylation.

3. Elongation of Polypeptide Chain:

As shown in Fig. 10.15 E, at the end of initiation sequence, the 70S ribosome possesses the fmet-tRNA in the P site, whereas the A site is free to receive the next aminoacyl-tRNA according to the codons on mRNA. The addition of amino acids to the growing polypeptide chain as per codon on mRNA is called elongation of chain.

The rate of addition of amino acid to the growing polypeptide is about 16 residues per second at 37°C. The 5S rRNA molecule recognises the nucleotide sequence of TѰ loop of tRNA and thus helps in binding of tRNA to the A site. The codons direct the specific aminoacyl-tRNA to form bonds. The bond formation is stimulated by an elongation factor T (EF- T) and GTP. T refers to transferase activity.

The elongation factor (EF) is a soluble protein which is required for elongation of polypeptide chain. The EF is of two types, EF-T and EF-G. The EF-T is associated with transferase activity, whereas the EF-G is involved in translocation of mRNA.

In prokaryotes the EF-T consists of two protein subunits which are called EF-Tu (temperature unstable, MW 44,000) and EF-Ts (temperature stable, MW 30,000). The EF-Tu is most abundant protein in E.coli that accounts for 5-10% of the total cellular protein. Both the proteins (EF-Tu and EF-Ts) are needed for binding the aminoacyl- tRNA to the ribosome.

The eukaryotic EF is called EF-1 and EF-2 which has resemblance with the prokaryotic EF-T and EF-G. More specifically the EF-1 is like the EF-Tu in its structure and function. At a time the EF-1 exists in one of the two forms (light form, EF- 1L and heavy form, EF- 1H).

The function of EF-2 is translocation of aminoacyl-tRNA from A site to the P site. The GTP is required to drive the process of chain elongation. Bermerk (1978) has discussed the mechanism of chain elongation on ribosome.

Elongation of the polypeptide chain is accomplished in the following two steps:

Fig. 10.16 : Events of polypeptide chain formation.

(i) Binding of Aminoacyl-tRNA to the A Site:

The GTP binds to EF-T and splits it into EF- Tu-GTP and EF-Ts. The EF-Tu-GTP can bind to all aminoacyl-tRNA (except the initiator tRNA) and results in formation of GTP-EF-Tu-aminoacyl-tRNA complex (Fig. 10.16A). It is an interme­diate complex which is bound to the ribosome.

In this step the EF-Ts complex does not play any role. After the aminoacyl-tRNA binds to the A site, GTP is hydrolysed and EF-Tu-GDP complex is released from the ribosome (B). Each aminoacyl -tRNA bound hydrolyses one GDP. The aminoacyl – tRNA may bind to the A site but this binding may not be followed by EF-Tu release from the ribosome. This shows that the purpose of GTP hydrolysis is the release of EF-Tu from the ribosome.

(ii) Peptide-Bond Formation:

Soon the enzyme peptidyl transferase (PTas) catalyses the peptide-bond formation. In fact this is catalysed by the 23S rRNA. This process is called peptidyl transfer (Fig. 10.16.C).

However, peptide bond formation depends on release of EF-Tu from the ribosome but not on hydrolysis of GTP. The EF-Ts complex recycles the EF-Tu-GDP to EF-Tu- GTP, but does not cause release of EF-Tu from the ribosome as the release of IF-2 depends on IF-1.

When a new aminoacyl-tRNA binds to the A site, peptide bond formation occurs between the starting amino acid (N-fmet-tRNA on prokaryotes and met-tRNA in eukaryotes) and new aminoacyl-tRNA at the P site.

The enzyme peptidyl transferase located in 50S ribosomal subunit catalyses the formation of peptide bond between the amino group of new incoming amino acid and the C-terminal of the elongating polypeptide attached to tRNA (Fig. 10.16D). During the process of bond formation, H2O is eliminated.

4. Translocation:

When the peptide bond is formed, the growing peptide chain binds to the tRNA that carries the incoming amino acid and occupies the A site of ribosome. The discharged tRNA after dissociating itself from the peptide chain is released from the P site (Fig. 10.16D-E). It is known so far that ribosome consists of two sites (A and P) but the recent evidences suggest that it consists of three sites: A, P and E. The E site is specific for de-acylated tRNA (E).

(i) Mechanism of Translocation:

In the ribosome at site A (aminoacyl-tRNA accepting site) the incoming aminoacyl-tRNA enters where decoding (codon-anticodon recognition) takes place. Thereafter, the ribosome moves along mRNA and, therefore, a change in complex occurs.

The movement of ribosome causes the alignment with A site of next codon of mRNA to be translated. Consequently, the peptidyl-tRNA situated at A site is transferred to P site. This event of transfer of peptidyl – tRNA is called translocation (Fig. 10.16E-F).

During translocation the events that are accomplished are:

(i) Removal of discharged tRNA from the P site,

(ii) Movement of the peptidyl tRNA from the A site to the P site, and

(iii) Movement of message by one codon.

(ii) Energetics:

The recent model of ribosome shows that:

(i) The incoming charged tRNA binds at the A site,

(ii) The growing polypeptide attached to tRNA and bound to P site is transferred to the tRNA in the A site, and

(iii) The newly deacaylated tRNA after translocation is not released immediately but are bound to the E site.

Now both the E and P sites are engaged. The other incoming charged tRNA binds to the unoccupied A site. This causes reduction in affinity of the E site for the deacylated tRNA and resulting in release of discharged tRNA from the ribosome. The process of binding of incoming aminoacyl-tRNA to site A continue until the termination signal is received (G-I).

In prokaryotes translocation is brought by the EF-G or translocase (MW, 80,000 in which G = GTPase) and GTP hydrolysis are required. EF-G binds to the same site as the EF-Tu. After binding EF-G hydrolyse the ATP to ADP + Pi in the presence of ribosome.

It is obvious that during elongation two molecules of GTP are hydrolysed per peptide bond, one is EF-T dependent and the other EF-G dependent. The EF-G is released from the ribosome after each step of elongation. Since both EF-T and EF-G utilize the same binding site, elongation cannot continue unless EF-G is released.

5. Termination of Polypeptide Chain:

(i) Recognition of Termination Signal:

The polypeptide chain continues the elongating until a termination codon on mRNA reaches to ribosome. The termination codons (UAA-ochre, UAG- amber, UGA-opal or umber) are also called as non-sense codon because no tRNA anticodon pairs with them. It is not necessary that the termination codon is the last codon of mRNA.

For example in bacteria and bacteriophages polygenic mRNAs are common and they consist of a number of initiation and termination codons. After translocation of one of the above termination codons into the A site, the ribosome does not bind to an aminoacyl-tRNA-EF-Tu-GTP complex. Then it receives the signal of termination.

(ii) Release of Polypeptide Chain:

When a termination codon is translocated into the A site, the ribosome instead of binding with a complex containing an amino acid, binds with a peptide release factor (RE) (Fig. 10.16 J). In prokaryotes there are three RF proteins (RF-1, MW 44,000 RF-2, MW 47,000 RF-3, MW 46,000).

The RF-1 is active with UAA and UAG codons and the RF-2 is active with UAA and UGA codons. The RF-3 activates the RF-1 and RF-2 therefore, the RF-3 is called stimulatory (S) factor. In eukaryotes there is only one RF protein (MW 56,500 and 1,15,000) which is active with codons UAA, UAG and UGA. The RF protein exists in two units and both of them remain in active form.

The ribosome binds either with RF-1 or RF-2. However, the RF protein activates peptidyl transferase which hydrolyses the bond joining the peptide to the tRNA at the site P. This results in release of the peptide chain (Fig. 10.16J).

6. Post Translational Processing:

After release some of the processing events occur in the polypeptide chain.

Such modifications occur both in prokaryotes and eukaryotes as given below:

(i) Removal of fmet from the Polypeptide Chain:

The formyl group of the N-terminal fmet is removed by the enzyme methionine deformylase. The enzyme formylmethionine specific peptidase (methionyl amino-peptidase or MAP) hydrolyses the entire formylmethionine residues. All the terminal methionine’s are not removed because there is involvement of discrimination in channeling of different polypeptides through these two alternative steps.

The side-chain penultimate amino acid acts as the discriminating factor. Removal of methionine by MAP depends on the length of side chain. The side chain of the longer length has less possibility for MAP to remove the methionine. The other processing’s are acetylation (of L12 to give rise L7) or adenylation.

(a) Loss of signal sequences:

In some polypeptides, about 15 to 30 amino acid residues are present at N-terminus. These residues act as signal sequence and direct the protein to its ultimate destination. The signal sequences are cleaved by specific peptidases.

(b) Modification of individual amino acid:

Some amino acid side chains are specifically modified such as:

(a) Enzymatic phosphorylation by ATP of -OH group of certain amino acids (e.g. serine, threonine, tyrosine),

(b) Binding of Ca ++ to phosphoresine groups of milk protein, casein,

(c) Addition of carboxyl (-COOH) group to aspartate glutamate residues of some proteins (e.g. blood clotting protein, prothrombin),

(d) Methylation of proteins (e.g. methylation of lysine residues in cytochrome c, calmodulin).

(c) Formation of disulphide cross-links:

Disulphide bridges between cysteine residues of some proteins are formed. Hence, they are covalently cross-linked and attain native from.

(d) Glycosylation:

Attachment of the carbohydrate side chains during or after protein synthesis is called glycosylation, for example glycoproteins.

(e) Addition of prosthetic group:

Prosthetic groups get covalently bound to many prokaryotic and eukaryotic proteins. For example, biotin molecule is covalently linked to acetyl-CoA carboxy­lase.

(ii) Ribosome Editing:

During the process of translation certain inappropriate amino-acylated – tRNAs enter the A site of ribosome and remain bound to the ribosome. These outnumber the appropriate amino acylated (aa)-tRNAs.

However, the aa-tRNAs remain bound for a long time to the A site for a peptide from P site of ribosome. There are two processes that can reduce the error of surviving polypeptide chain e.g. ribosome editing and preferential degradation of polypeptide chain containing erroneous amino acids.

According to the ribosome editing hypothesis the structure of inappropriate peptidyl-tRNA does not correctly complement the structure of mRNA therefore, it dissociates from the ribosome during protein synthesis. However, the gene relA produces a signal molecule (alarmone), guanosine tetra-phosphate (ppGpp) which affects the editing process.

The ppGpp interacts with EF-G and results in longer life of peptidyl-tRNA in the A site and en-chances the editing process. The gene pth synthesizes peptidyl-tRNA hydrolase that acts upon the peptidyl-tRNA when it is released. The ribosome free peptidyl-tRNA is hydrolysed by this enzyme. Consequently an intact peptide and an intact tRNA are produced. The defective peptide is degraded by the enzyme.

(iii) Protein Folding:

After the synthesis of polypeptide chain, it undergoes spontaneous foldings. The secondary folds are formed between the folded regions. Finally as a result of further folding there develops a tertiary structure of polypeptide chain i.e. protein. Before the terminal amino acids are added in polypeptide chain, the protein reaches to its final shape during the course of chain termination.

According to the recent theories the process of protein folding is complex. Many proteins require assistance to get folded properly. This assistance is provided by the proteins of special kind known as chaperones or chaperonins. These assist polypeptides to self assemble by inhibiting alternative assembling pathway.

When chaperones interact with the polypeptide, the chance of incorrect in-folding is reduced. In E. coli the example of chaperones are GroE1 (60 KDa), GroES (10KDa) and DnaK (70KDa). All these are constitutive proteins which increase their concentration when there is stress like heat shock.

7. The Signal Hypothesis (Protein Export):

It is interesting to note that after synthesis of protein, how it is incorporated in membranes or secreted outside by the cell? However, it is believed that the secretory proteins are synthesised by the ribosomes which are attached to the endoplasmic reticulum and released into it. From endoplasmic reticulum they are transported to various cell organelles (in eukaryotes) where from secreted outside the cell through the process of exocytosis.

To explain this mechanism Blobel and Dobberstein (1975) proposed a theory known as signal hypothesis. Furthermore Blobel (1978) reviewed this hypothesis and postulated that the mRNAs that translate secretory proteins possess on 3′ side of initiation codon (AUG) a group of signal codons.

The endoplasmic reticulum consists of ribosome receptor proteins. A polypeptide chain consisting of a special region (signal peptide region) is synthesized by the ribosome. After coming out from the ribosome the signal peptide interacts with ribosome receptor protein and results in formation of a tunnel in the membrane.

However, the membrane tunnel coincides with the ribosomal tunnel. The enzyme signal peptidase breaks the polypeptide chain which is being synthesised. Upon complete synthesis of polypeptide chain, it is released inside the space of endoplasmic reticulum.

At the end the ribosomes dissociate from the membrane of endoplasmic reticulum ribosome receptor proteins get diffused and close the tunnels. This process of entering the proteins into the membranes is also known as protein export.

A significant work has been done in recent years on secretory protein translocation system in E. coli and the other bacteria as well such as Bacillus subtilis. Salmonella typhimurium, Pseudomonas fluorescens, Enterobacter aero genes. Vibrio cholerae, Klebsiella oxycota, etc.

Most of the secretory proteins are translated first in a form of precursor containing 15-30 amino acids at N-terminus which is called a signal sequence. The signal sequence consists of a hydrophobic region of about 11 amino acid residues and a short stretch of hydrophilic region at N-terminus. The signal sequence is involved to bind the nascent polypeptide to the membrane.

In Gram-negative organisms the pathway for export and secretion of signal sequence-containing proteins is called general secretory pathway. The first step is the sec gene product depen­dent translocation of exported protein outside the cytoplasmic membrane. Brondage (1990) have presented a model of proOmpA (an export pro­tein) translocation across the plasma membrane (Fig. 10.17).

Fig. 10.17 : A model for translocation of proOmpA across the plasma membrane.

The protein SecB (a product of secB gene) is a pilot chaperonin. It is associated with the protein that is to be transported i.e. transport protein, for example proOmpA. It is also synthesised on the ribosome. However, in the absence of SecB, the proOmpA aggregates and checks its insertion into the membrane.

The protein SecA is basically an ATPase and also forms a part of pre-protein translocase in association with integral membrane protein SecY/E. The SecA protein acts as a receptor for proOmpA-SecB complex. Subse­quently, the ATP is hydrolysed releas­ing the proOmpA into the membrane. It also drives the overall chaperones and membrane-associated reactions.

Once the process of transport has been started at the expense of ATP, further translocation event proceeds through a series of trans-membrane intermediates, the energy requirement of which is met by proton motive force rather than ATP hydrolysis. During the process of translocation, the enzyme signal peptidase (LepB) cleaves the signal sequence of the exported protein.

It has to enter the periplasm or translocate across the outer membrane. Pugsley (1993) has published the complete general secretary pathway in Gram-negative bacteria where the various branches direct proteins to their final extra cytoplasmic destination (Fig. 10.18).

Fig. 10.18 : Main branches of the general secretory pathways (GSP) of Gram-negative bacteria. IMP, integrated membrane proteins CAC, chaperone assembly channel PSI, periplasmic secrection intermediates SP, signal peptidase.

8. The Inhibitors of Gene Expression:

There are several antimicrobial agents that inhibit protein synthesis at two steps, either during transcription or translation. Franklin and Snow (1991) have nicely discussed the biochem­istry of the action of antimicrobial antibiotics.

(i) Inhibitors of Transcription:

The rifamycin and chemically similar group (streptovarcins) inhibit the initiation of transcription. Rifampin and streptovarcin are the semisynthetic compounds, whereas rifamycins are the naturally occurring antibiotics. They tightly bind to P subunit of RNA polymerase (rpoB) and inhibit initiation of transcription.

(b) Streptolydigin:

It is similar to rifamycin because it too binds with P subunit of RNA polymerase. In addition, it inhibits both the processes: chain initiation and chain elongation in vitro.

(ii) Inhibitors of Translation:

It inhibits the activity of peptidyl transferase after binding to 508 subunit of bacterial ribosome. Its effect is bacteriostatic i.e. after removal of drug the effect is soon reversed. However, its effect in eukaryotes is the same as in prokaryotes. Bone marrow toxicity results in aplastic anaemia.

This group of antibiotic shows broad spectrum bacteriostatic activities against Gram-positive and negative bacteria, mycoplasmas, rickettsiae and chlamydiae. These prevent the binding of aminoacyl -tRNA to the A site of 30S ribosome. Moreover, it can bind to several sites of both 30S and 50S subunits.

(c) Cycloheximide (actidione):

It inhibits protein synthesis in eukaryotes (e.g. yeasts, fungi, higher plants and mammals) but not the prokaryotic microorganisms. It interferes with the activity of ribosome present in cytoplasm but not in mitochondria, by binding with 80S subunits and preventing the movement of mRNA.

This is a large group of antimicrobial agents that includes erythromycin, leucomycin, macrocin, carbomycin, chalcomycin, angalomycin, etc. These are active against the Gram-positive bacteria and less active against Gram-negative bacteria but not against the eukaryotes. These interact with SOS subunit of ribosome and inhibit protein synthesis. Also, they stimulate the dissociation of peptidyl-tRNA from the ribosome through abortive translocation step.

Lincomycin and the other chemically similar antibiotics inhibit peptidyl transferase through binding to 23S rRNA of the 50S subunit. These affect both Gram-positive and Gram-negative bacteria.

It binds to a peptide with the C-terminus of growing polypeptide and results in premature termination of polypeptide chain. It interacts with the P site of ribosome but not A site. It works equally well on 70S and 50S ribosomes.


Introduction

Following synthesis by RNA polymerase II, eukaryotic mRNA precursors (pre-mRNA) undergo a series of modifications, including capping at the 5′ end, splicing and addition of a 3′ poly(A) tail. The poly(A) tail has been shown to affect virtually all aspects of mRNA metabolism, including translation initiation and mRNA stabilization (reviewed in Beelman and Parker, 1995 Sachs et al., 1997 Wahle and Rüegsegger, 1999 ). Polyadenylation is a two-step event: endonucleolytic cleavage of the pre-mRNA is followed by poly(A) addition to the 3′ end of mRNAs. The cleavage and polyadenylation reactions involve a multicomponent machinery of remarkable complexity (reviewed in Colgan and Manley, 1997 Wahle and Rüegsegger, 1999 Zhao et al., 1999 ). In mammals, the synthesis of the poly(A) tail is catalyzed by poly(A) polymerase (PAP) with the help of the cleavage and polyadenylation specificity factor (CPSF Christofori and Keller, 1988 Takagaki et al., 1988 Gilmartin and Nevins, 1989 Wahle, 1991b ). CPSF binds to the canonical polyadenylation signal AAUAAA, which is located in the pre-mRNA 10–30 nucleotides upstream of the polyadenylation site ( Keller et al., 1991 ). PAP alone binds RNA very weakly and non-specifically and shows a low, distributive polyadenylation activity. This is in contrast to what happens after the polymerase is recruited to AAUAAA-containing RNAs by CPSF the protein–protein interactions with CPSF stimulate polyadenylation and greatly stabilize the ternary complex composed of CPSF, PAP and the RNA primer ( Bienroth et al., 1993 Murthy and Manley, 1995 ). In the presence of the nuclear poly(A)-binding protein II (PABP2), another factor of the polyadenylation machinery, poly(A) synthesis becomes fast and processive. PABP2 has also been shown to control the final length of poly(A) tails, which on average are 250 nucleotides long in vivo ( Wahle, 1991a Wahle et al., 1993 ).

Most PAPs are single polypeptide enzymes, and cDNA clones encoding them have been isolated from many different organisms ( Martin et al., 1999 ). PAPs have a modular organization with a catalytic domain near the N-terminus and an RNA-binding region that overlaps with a nuclear localization signal (NLS) near the C-terminus ( Zhelkovsky et al., 1995 Martin and Keller, 1996 Martin et al., 1999 ). C-terminal extensions of ∼20 kDa are only found in vertebrates and are dispensable for catalytic activity in vitro. The extended C-terminal domain of vertebrate PAPs is rich in serines and threonines, and enzyme activity can be downregulated by phosphorylation at multiple sites (reviewed in Colgan and Manley, 1997 Wahle and Rüegsegger, 1999 ). The extreme C-terminus of PAP is also the target for another type of regulation. The U1A protein, a component of the U1 snRNP which functions in 5′ splice site recognition, is known to inhibit polyadenylation of its own mRNA by binding to PAP ( Gunderson et al., 1994 Vagner et al., 2000 ). The C-terminus of PAP is also involved in protein–protein interactions with the splicing factor U2AF65 ( Vagner et al., 2000 ) and the snRNP protein U1-70K ( Gunderson et al., 1998 ).

PAPs belong to a large superfamily of nucleotidyl transferases containing two signature features: a conserved catalytic domain adorned with three invariant carboxylates that are crucial for activity, and a helical turn motif involved in nucleotide binding ( Holm and Sander, 1995 Martin and Keller, 1996 Aravind and Koonin, 1999 Martin et al., 1999 ). Mammalian DNA polymerase β (pol β) and kanamycin nucleotidyl transferase (KanNt) are also members of this superfamily of nucleotidyl transferases. The structures of pol β ( Davies et al., 1994 Pelletier et al., 1994 Sawaya et al., 1997 ) and KanNt ( Sakon et al., 1993 Pedersen et al., 1995 ) are known. Both crystal structures have shown that conserved residues in the helical turn motif are involved in recognizing the triphosphate moiety of the nucleotide. Pol β is a template-dependent DNA polymerase involved in the base excision repair pathway ( Pelletier et al., 1994 Sobol et al., 1996 Sawaya et al., 1997 ). Until now, a structure of a template-independent polymerase such as PAP was lacking.

Here we report the structure of a C-terminal deletion mutant of bovine PAP bound to an analog of ATP, cordycepin 5′-triphosphate (3′-dATP), at 2.5 Å resolution. The structural results presented here reveal a surprising domain composition and architecture, and provide insight into the ways in which PAP specifically binds to and incorporates ATP.


Major concerns with the integrity of the mitochondrial ADP/ATP carrier in dodecyl-phosphocholine used for solution NMR studies

Membrane proteins (MPs) comprise 15-25% of the human proteome, yet collectively sum to less than 1% of the deposited three-dimensional structures in the Protein DataBank (PDB). Such a paucity of structural information manifests from difficulties that can accompany the biochemical preparation of purified MPs, ranging from complications with faithful abstraction from the native membrane to misfolding after extraction from insoluble inclusion bodies. Moreover, despite the common view of MPs existing in one of two end states (i.e. “open” or “closed” transporters, “active” or “inactive” G-protein coupled receptors), MPs can populate a wide range of conformations that dynamically exchange on various timescales, spanning pico-nanoseconds, micro-milliseconds, or far longer than hours. Such dynamics can often hinder crystallization attempts for structural analysis by X-ray crystallography. Despite the current lack of structural knowledge pertaining to MPs, many FDA-approved drugs target them. Thus, a holistic understanding of membrane protein structures, dynamics, and interactions in native membrane environments, or reliable membrane mimetics, proves critical.

A significant factor in structural studies of MPs is the membrane environment. As pointed out in a recent review 1 , nearly 80% of MP structures determined to date have used detergents, which form small micelles that present a very different environment to lipids. However, recent work has revealed that many MPs embedded in such micelles, for instance of the commonly used detergent dodecyl phosphocholine (DPC, also known as Foscholine-12), are no longer functional or folded correctly 1 .

Thus, when choosing a membrane mimetic system, how does one ensure that a reconstituted MP is functionally relevant? The most rigorous demonstration is an assay that directly monitors biochemical activity: for example, the transport of a substrate or other reporter molecule. In the absence of such an assay, researchers typically default to the demonstration of the MP binding to a relevant ligand, generally by measuring a dissociation constant (Kd) and comparing this to values obtained by other groups who have used native membranes. In the preprints discussed below, two reproduction studies are conducted pointing out significant flaws in this practice, and show that the presumed functionality of MPs should be interpreted with caution when using detergents.

The preprints

Two recent preprints conducted reproducibility studies on the structural and functional integrity of an AAC transporter (AAC3) in DPC 2,3 . AAC3 functions to exchange ADP for ATP across the inner mitochondrial membrane, an essential process for the cell. This membrane protein had previously been crystallized in the presence of a strong inhibitor (CATR) and was shown to adopt a locked conformation, termed the “c-state” 4,5 . As part of its transport mechanism, AAC3 populates a second conformation known as the “m-state”, yet structural evidence for this state has remained elusive.

In 2015, a paper by Chou and colleagues 6 in Nature Structural & Molecular Biology reported using NMR spectroscopy that, in DPC, the yeast version of AAC3 (yAAC3) could be captured in a dynamic equilibrium involving two states: the c-state and a transiently populated conformation, which was inferred to be the m-state (or similar). Critically, their results hinged on yAAC3 remaining correctly folded and fully functional while embedded in the detergent. The integrity and correct folding of the yAAC3 sample was determined by binding of the inhibitor (CATR) and substrate (ADP) as measured by isothermal titration calorimetry (ITC) or NMR 6 .

Preprints from Schanda and colleagues 2 at the Institut de Biologie Structurale in Grenoble (France) and Kunji and coworkers 3 at the Mitochondrial Biology Unit in Cambridge (UK) challenged the notion that yAAC3 is correctly folded and functional in DPC. Rather, the authors convincingly demonstrate that yAAC3 is misfolded and not functional in the presence of DPC.

First, both King et al. 3 and Kurauskas et al. 2 noted that the Kd values for CATR binding to yAAC3 reported by Chou and colleagues were nearly three-to-four orders of magnitude larger than the commonly accepted literature values (5–300 nM). Second, Schanda and colleagues examined the raw data used to determine that the substrate (ADP) binds specifically, and found residues all over yAAC3, including those more than 20 Å distant from the ADP binding site, were impacted by ADP binding 2 . While this could reflect allosteric structural changes, a recent publication noted that AAC3 embedded in DPC bound identically to ATP and GTP, despite preference for the former in the native membrane 5 . Such a loss of specificity implies an incorrect tertiary structure. Finally, King et al. 3 measured melting temperatures of yAAC3 in DPC and other detergent systems (Figure 1A). In the native mitochondrial membrane, yAAC3 unfolded near 40°C however, the same experiment performed in DPC revealed a large fraction of unfolded protein already present at ambient temperature, which exhibited no unfolding cooperativity 3 . Similar results were obtained by NMR 3 . Taken together, these data suggest that yAAC3 solubilized in DPC is incorrectly folded and binds to CATR and ADP non-specifically 2,3 . King et al. note that a non-specific interaction could be electrostatic in nature, as yAAC3 has an isoelectric point of 9.82, and therefore contains excess positive charge, whereas ADP and CATR are both negatively charged at neutral pH 3 .

Having established that ADP and CATR binding in fact reflects non-specific interactions, Schanda and colleagues 2 re-examined the previously published NMR data used to measure conformational exchange in yAAC3 between the c-state and putative m-state (Figure 1B). The experiment used for such analysis is called Carr-Purcell-Meiboom-Gill relaxtion dispersion (CPMG RD, reviewed here), and involves the measurement of an NMR parameter (transverse relaxation rate, termed the effective R2) as a function of the number of radiofrequency pulses applied during a fixed delay. In the presence of conformational exchange between two or more states on the micro-to-millisecond timescale, CPMG RD experiments will reveal a dependence of R2 on the number of applied radiofrequency pulses (Figure 1B). Subsequently fitting these data to the appropriate equations yields the rate of conformational exchange, the relative populations, and (qualitative) residue-specific information about structural differences between the two forms. For example, such experiments have been deployed to determine accurate solution structures of otherwise invisible protein folding intermediates that exchange with a folded conformation on the millisecond timescale 7 .

In the original paper, Chou and colleagues 6 found via CPMG RD that the rate of conformational exchange varied by roughly 20-fold between the inhibitor- and substrate-bound forms of yAAC3, and reported large structural differences between the two states (for NMR spectroscopists, 15 N |delta omega| > 5 ppm). However, Kurauskas et al. re-fit the raw CPMG RD data in free state and the inhibitor- and substrate-bound forms of yAAC3 (Figure 1B), finding that no difference existed in the rate of conformational exchange (1500 s -1 for all forms), with significantly smaller structural differences than initially reported ( 15 N |delta omega| values around 2 ppm). Finally, similar millisecond exchange dynamics that were independent of added substrate/inhibitor were obtained when analyzing conformational exchange in mutants of AAC3 that are incapable of ADP/ATP exchange, indicating that there is no link between dynamics and transport 8 .

In conclusion, these two new preprints from the Schanda and Kunji groups demonstrate that the yAAC3 membrane protein is not functional and misfolded when embedded in DPC. These studies highlight the difficulty and importance of choosing the right membrane mimetic (when not using native membranes). In addition, these works provide a platform for diagnosing future issues that arise from misfolded MPs (e.g. non-specific ligand binding, lower Kd than expected, pervasive millisecond motions), in particular those analyzed by NMR spectroscopy.

These two new preprints highlight the importance of independent replication studies that seek to verify, challenge, or question biological implications obtained from biophysical experiments. While a few examples exist in the published literature, there are certainly not enough of such studies. Encouragingly, a replication preprint was recently highlighted on preLights. The bioRxiv will help to remove the stigma associated with publishing replication studies, and will provide a convenient platform for the publication of replication studies in lieu of publication in traditional journals. Likewise, PLOS Biology has instituted an initiative to publish papers from groups who have recently been “scooped”, which should also increase the number of papers that report data on the same system.

While the two preprints discussed here present negative results, their findings will together advance the field of AAC structural biology, and hopefully will deter future researchers from using DPC as a reconstitution system. More generally, the preprints from the Schanda and Kunji groups highlight the importance of selecting the appropriate membrane mimetic system and then verifying that the embedded MP retains functional and structural integrity.

As a practicing NMR spectroscopist who enjoys dynamical measurements, the experiment that I most enjoyed was actually a data analysis method to obtain insight into micro-to-millisecond motions. Schanda and colleagues noted that the CPMG RD data from Chou and coworkers could be described by a single set of parameters for all three conditions (apo-AAC3, CATR-bound AAC3, ADP-bound AAC3), and showed that these data could indeed be fitted globally. This means that the AAC3 exchange process on the micro-to-millisecond timescale, which is probed in the NMR experiment, is insensitive to added substrate or inhibitor, and is therefore not a transport filter.


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