Enzymatic Conversion of Glucose to UDP-4-Keto-6-Deoxyglucose in Streptomyces spp.

All of the 2,6-dideoxy sugars contained within the structure of chromomycin A₃ are derived from d-glucose. Enzyme assays were used to confirm the presence of hexokinase, phosphoglucomutase, UDPG pyrophosphorylase (UDPGP), and UDPG oxidoreductase (UDPGO), all of which are involved in the pathway of glucose activation and conversion into 2,6-dideoxyhexoses during chromomycin biosynthesis. Levels of the four enzymes in Streptomyces spp. cell extracts were correlated with the production of chromomycins. The pathway of sugar activation in Streptomyces spp. involves glucose 6-phosphorylation by hexokinase, isomerization to G-1-P catalyzed by phosphoglucomutase, synthesis of UDPG catalyzed by UDPGP, and formation of UDP-4-keto-6-deoxyglucose by UDPGO.Dideoxy sugars occur commonly in the structures of cardiac glycosides from plants, in antibiotics like chromomycin A₃ (Fig. ​(Fig.1),1), and in macrolides produced by microorganisms. On the basis of stable isotope-labeling experiments, biosynthetic studies conducted in Rosazza’s laboratory have indicated that all the deoxy sugars of chromomycin A₃ are derived from d-glucose (21). While the assembly of the polyketide aglycone is reasonably well understood, relatively little is known of the details of 2,6-dideoxy sugar biogenesis in streptomycetes. Earlier studies with Streptomyces rimosus indicated that TDP-mycarose is synthesized from TDP-d-glucose (TDPG) and S-adenosyl-l-methionine (10, 23). The reaction requires NADPH as a cofactor, and TDP-4-keto-6-deoxy-d-glucose is an intermediate. Formation of TDP-4-keto-6-deoxy-d-glucose was catalyzed by the enzyme TDPG oxidoreductase (TDPG-4,6-dehydratase; EC 4.2.1.46). Similar 4-keto sugar nucleotides are intermediates for the biosynthesis of polyene macrolide antibiotic amino sugars (18). Similar pathways have been elaborated for the formation of 2,6-dideoxy-d-threo-4-hexulose of granaticin in Escherichia coli (6, 25) and 2,6-dideoxy-d-arabino-hexose of chlorothricin (12). The initial 6-deoxygenation of glucose during 3,6-dideoxy sugar formation involves a similar mechanism (32). In all of these processes, glucose is first activated by conversion into a sugar nucleotide such as UDPG followed by NAD⁺ oxidation of the 4 position to the corresponding 4-oxo derivative. Position 6 deoxygenation involves a general tautomerization, dehydration, and NADH,H⁺-catalyzed reduction process (6, 12, 25). A similar tautomerization and dehydration followed by reduction may produce C-3-deoxygenated products, such as CDP-3,6-dideoxyglucose (27). The pathway for formation of 3,6-dideoxyhexoses from CDPG in Yersinia pseudotuberculosis was clearly elucidated by Liu and Thorson (14). However, none of this elegant work was focused on the earlier steps of hexose nucleotide formation. Open in a separate windowFIG. 1Structures of chromomycins A₂ and A₃.On the basis of previous work (7), it is reasonable to postulate that the biosynthesis of 2,6-dideoxyglucose in Streptomyces griseus involves phosphorylation to glucose-6-phosphate by hexokinase (HK; E.C.2.7.7.1), as in glycolysis; conversion to glucose-1-phosphate by phosphoglucomutase (PGM; EC 2.7.5.1); reaction with UTP to form UDPG in a reaction catalyzed by UDPG pyrophosphorylase (UDPGP) (glucose-1-phosphate uridylyltransferase; EC 2.7.7.9), and C-6 deoxygenation catalyzed by UDP-d-glucose-4,6-dehydratase with NAD⁺ as a cofactor (Fig. ​(Fig.2).2). UDPG and GDPG have been detected in cell extracts of S. griseus and Streptomyces sp. strain MRS202, suggesting that these compounds are active sugar nucleotides involved in the formation of dideoxyhexoses (15). UDPGP genes from several bacteria have been cloned and sequenced (1, 3, 4, 11, 29, 30). Although nucleotidyl diphosphohexose-4,6-dehydratases (NDP-hexose-4,6-dehydratases) have been purified and characterized from several sources (5, 8, 9, 13, 19, 25, 26, 31, 33), the occurrence of the glucose-activating enzymes HK, PGM, UDPGP, and UDPG oxidoreductase (UDPGO) involved in 2,6-dideoxyhexose formation has not been established in streptomycetes. This work provides evidence for the presence of these enzymes involved in the biosynthetic activation of glucose to the 2,6-dideoxyhexoses in chromomycin A₃.Open in a separate windowFIG. 2Proposed pathway for the formation of 2,6-dideoxy sugars in streptomycetes involving HK, PGM, UDPGP, and UDPGO.     

Resistance of Human Cytomegalovirus to Benzimidazole Ribonucleosides Maps to Two Open Reading Frames: UL89 and UL56

2,5,6-Trichloro-1-β-d-ribofuranosyl benzimidazole (TCRB) is a potent and selective inhibitor of human cytomegalovirus (HCMV) replication. TCRB acts via a novel mechanism involving inhibition of viral DNA processing and packaging. Resistance to the 2-bromo analog (BDCRB) has been mapped to the UL89 open reading frame (ORF), and this gene product was proposed as the viral target of the benzimidazole nucleosides. In this study, we report the independent isolation of virus that is 20- to 30-fold resistant to TCRB (isolate C4) and the characterization of the virus. The six ORFs known to be essential for viral DNA cleavage and packaging (UL51, UL52, UL56, UL77, UL89, and UL104) were sequenced from wild-type HCMV, strain Towne, and from isolate C4. Mutations were identified in UL89 (D344E) and in UL56 (Q204R). The mutation in UL89 was identical to that previously reported for virus resistant to BDCRB, but the mutation in UL56 is novel. Marker transfer analysis demonstrated that each of these mutations individually caused ∼10-fold resistance to the benzimidazoles and that the combination of both mutations caused ∼30-fold resistance. The rate and extent of replication of the mutants was the same as for wild-type virus, but the viruses were less sensitive to inhibition of DNA cleavage by TCRB. Mapping of resistance to UL56 supports and extends recent work showing that UL56 codes for a packaging motif binding protein which also has specific nuclease activity (E. Bogner et al., J. Virol. 72:2259–2264, 1998). Resistance which maps to two different genes suggests that their putative proteins interact and/or that either or both have a benzimidazole ribonucleoside binding site. The results also suggest that the gene products of UL89 and UL56 may be antiviral drug targets.Human cytomegalovirus (HCMV) can cause significant morbidity and mortality in immunocompromised populations (3). It is a common opportunistic disease in patients with AIDS and is often a factor in their death (38). HCMV infection has been implicated in increased risk of organ rejection following heart (28) and kidney transplants (8) and in restenosis of diseased arteries following angioplasty (41, 63). It is also a leading cause of birth defects (16).Current therapies for HCMV infection include ganciclovir (GCV) (22), cidofovir (30), and foscarnet (20). Each of these drugs has several limitations to its use: none are orally bioavailable, all have dose-limiting toxicity, and resistance has developed to each (26). Because all three of these drugs inhibit viral replication through an interaction with the virally encoded DNA polymerase (25, 31, 37), the possibility of cross-resistance exists. Thus, additional drugs with unique mechanisms of action are needed for the treatment of HCMV infections.In 1995, we reported that 2-bromo-5,6-dichloro-1-(β-d-ribofuranosyl)benzimidazole (BDCRB; Fig. ​Fig.1)1) and the 2-chloro analog [2,5,6-trichloro-1-(β-d-ribofuranosyl)benzimidazole TCRB] are potent and selective inhibitors of HCMV replication (55). These compounds have a novel mechanism of action, which unlike the current therapies for HCMV infection, does not involve inhibition of DNA synthesis. The benzimidazole ribonucleosides prevent the cleavage of high-molecular-weight viral DNA concatemers to monomeric genomic lengths (57). Resistance to BDCRB has been mapped to the HCMV UL89 open reading frame (ORF), which, by analogy to gene gp17 from bacteriophage T4, may be a terminase (23, 57). Consequently, we have proposed that the benzimidazole ribonucleosides inhibit the product of this gene and that the UL89 gene product is involved in the viral DNA concatemer cleavage process (57). Open in a separate windowFIG. 1Structure of benzimidazole ribonucleosides. TCRB, R = Cl; BDCRB, R = Br.HCMV replication proceeds in a manner which is conserved among herpesviruses. The virally encoded DNA polymerase produces large, complex head-to-tail concatemers (10, 29, 33) which must be cleaved into genomic-length pieces before insertion into preformed capsids (59). With herpes simplex virus type 1 (HSV-1), temperature-sensitive mutants which are unable to cleave and package the concatemeric DNA have been derived (1, 2, 4, 45, 49, 50, 61). By this process, six HSV-1 genes have been found to be involved in concatemer cleavage and packaging. They are UL6, UL15, UL25, UL28, UL32, and UL33. In addition, recent studies in Homa’s laboratory have established that the product of UL25 is required for viral DNA encapsidation but not cleavage (39). Homologs of these genes exist in HCMV and are UL104, UL89, UL77, UL56, UL52, and UL51, respectively (18).In our continuing investigation of the mode of action of benzimidazole nucleosides, we report herein the independent isolation of HCMV strains resistant to TCRB, characterization of these strains, and identification of the mutations responsible for the development of resistance. The results demonstrate that the mechanism of action of the benzimidazole ribonucleosides is more complex than previously proposed and that a second gene product implicated in DNA cleavage and packaging is involved.     

Limonene-1,2-Epoxide Hydrolase from Rhodococcus erythropolis DCL14 Belongs to a Novel Class of Epoxide Hydrolases

An epoxide hydrolase from Rhodococcus erythropolis DCL14 catalyzes the hydrolysis of limonene-1,2-epoxide to limonene-1,2-diol. The enzyme is induced when R. erythropolis is grown on monoterpenes, reflecting its role in the limonene degradation pathway of this microorganism. Limonene-1,2-epoxide hydrolase was purified to homogeneity. It is a monomeric cytoplasmic enzyme of 17 kDa, and its N-terminal amino acid sequence was determined. No cofactor was required for activity of this colorless enzyme. Maximal enzyme activity was measured at pH 7 and 50°C. None of the tested inhibitors or metal ions inhibited limonene-1,2-epoxide hydrolase activity. Limonene-1,2-epoxide hydrolase has a narrow substrate range. Of the compounds tested, only limonene-1,2-epoxide, 1-methylcyclohexene oxide, cyclohexene oxide, and indene oxide were substrates. This report shows that limonene-1,2-epoxide hydrolase belongs to a new class of epoxide hydrolases based on (i) its low molecular mass, (ii) the absence of any significant homology between the partial amino acid sequence of limonene-1,2-epoxide hydrolase and amino acid sequences of known epoxide hydrolases, (iii) its pH profile, and (iv) the inability of 2-bromo-4′-nitroacetophenone, diethylpyrocarbonate, 4-fluorochalcone oxide, and 1,10-phenanthroline to inhibit limonene-1,2-epoxide hydrolase activity.Epoxides are highly reactive compounds which readily react with numerous biological compounds, including proteins and nucleic acids. Consequently, epoxides are cytotoxic, mutagenic, and potentially carcinogenic, and there is considerable interest in biological degradation mechanisms for these compounds.In bacteria, epoxides are formed during the metabolism of alkenes (23) and halohydrins (15, 26, 34, 49). Enzymes belonging to a large number of enzyme classes, including dehydrogenases (17), lyases (21), carboxylases (1, 43), glutathione S-transferases (6, 8), isomerases (24), and hydrolases (7, 19, 44), are involved in the microbial degradation of epoxides.Epoxide hydrolases are enzymes catalyzing the addition of water to epoxides forming the corresponding diol. This group of enzymes has been extensively studied in mammals, while only limited information is available on bacterial epoxide hydrolases. Three functions for epoxide hydrolases are recognized (42). In bacteria, epoxide hydrolases are involved in the degradation of several hydrocarbons, including 1,3-dihalo-2-propanol (34), 2,3-dihalo-1-propanol (15, 26), epichlorohydrin (46), propylene oxide (16), 9,10-epoxy fatty acids (30, 36), trans-2,3-epoxysuccinate (2), and cyclohexene oxide (14). Other epoxide hydrolases, such as microsomal and cytosolic epoxide hydrolase from mammals (for reviews, see references 4, 8, and 44), are involved in the detoxification of epoxides formed due to the action of P-450-dependent monooxygenases (8). Epoxide hydrolases are also involved in biosynthesis of hormones, such as leukotrienes and juvenile hormone (40, 45), and plant cuticular elements (11). Remarkably, the bacterial and eukaryotic epoxide hydrolases described so far form a homogeneous group of enzymes belonging to the α/β-hydrolase fold superfamily (10, 38).Rhodococcus erythropolis DCL14, a gram-positive bacterium, is able to grow on both (+)- and (−)-limonene as the sole source of carbon and energy (47). Cells grown on limonene contained a novel epoxide hydrolase that does not belong to the α/β-hydrolase fold superfamily. This limonene-1,2-epoxide hydrolase converts limonene-1,2-epoxide to limonene-1,2-diol (p-menth-8-ene-1,2-diol [Fig. 1]). In this report, we describe the purification and characterization of this enzyme and show that limonene-1,2-epoxide hydrolase belongs to a novel class of epoxide hydrolases. Open in a separate windowFIG. 1Reaction catalyzed by limonene-1,2-epoxide hydrolase.     

Genetic Analysis of the nuo Locus,Which Encodes the Proton-Translocating NADH Dehydrogenase in Escherichia coli

Complex I (EC 1.6.99.3) of the bacterium Escherichia coli is considered to be the minimal form of the type I NADH dehydrogenase, the first enzyme complex in the respiratory chain. Because of its small size and relative simplicity, the E. coli enzyme has become a model used to identify and characterize the mechanism(s) by which cells regulate the synthesis and assembly of this large respiratory complex. To begin dissecting the processes by which E. coli cells regulate the expression of nuo and the assembly of complex I, we undertook a genetic analysis of the nuo locus, which encodes the 14 Nuo subunits comprising E. coli complex I. Here we present the results of studies, performed on an isogenic collection of nuo mutants, that focus on the physiological, biochemical, and molecular consequences caused by the lack of or defects in several Nuo subunits. In particular, we present evidence that NuoG, a peripheral subunit, is essential for complex I function and that it plays a role in the regulation of nuo expression and/or the assembly of complex I.

Complex I (NADH:ubiquinone oxidoreductase; EC 1.6.99.3), a type I NADH dehydrogenase that couples the oxidation of NADH to the generation of a proton motive force, is the first enzyme complex of the respiratory chain (2, 35, 47). The Escherichia coli enzyme, considered to be the minimal form of complex I, consists of 14 subunits instead of the 40 to 50 subunits associated with the homologous eukaryotic mitochondrial enzyme (17, 29, 30, 48–50). E. coli also possesses a second NADH dehydrogenase, NDH-II, which does not generate a proton motive force (31). E. coli complex I resembles eukaryotic complex I in many ways (16, 17, 30, 49): it performs the same enzymatic reaction and is sensitive to a number of the same inhibitors, it consists of subunits homologous to those found in all proton-translocating NADH:ubiquinone oxidoreductases studied thus far, it is comprised of a large number of subunits relative to the number that comprise other respiratory enzymes, and it contains flavin mononucleotide and FeS center prosthetic groups. Additionally, it possesses an L-shaped topology (14, 22) like that of its Neurospora crassa homolog (27), and it consists of distinct fragments or subcomplexes. Whereas eukaryotic complex I can be dissected into a peripheral arm and a membrane arm, the E. coli enzyme consists of three subcomplexes referred to as the peripheral, connecting, and membrane fragments (29) (Fig. ​(Fig.1A).1A). The subunit composition of these three fragments correlates approximately with the organization of the 14 structural genes (nuoA to nuoN) (49) of the nuo (for NADH:ubiquinone oxidoreductase) locus (Fig. ​(Fig.1B),1B), an organization that is conserved in several other bacteria, including Salmonella typhimurium (3), Paracoccus denitrificans (53), Rhodobacter capsulatus (12), and Thermus thermophilus (54). The 5′ half of the locus contains a promoter (nuoP), previously identified and located upstream of nuoA (8, 49), and the majority of genes that encode subunits homologous to the nucleus-encoded subunits of eukaryotic complex I and to subunits of the Alcaligenes eutrophus NAD-reducing hydrogenase (17, 29, 30, 49). In contrast, the 3′ half contains the majority of the genes that encode subunits homologous to the mitochondrion-encoded subunits of eukaryotic complex I and to subunits of the E. coli formate-hydrogen lyase complex (17, 29, 30, 49). Whereas the nuclear homologs NuoE, NuoF, and NuoG constitute the peripheral fragment (also referred to as the NADH dehydrogenase fragment [NDF]), the nuclear homologs NuoB, NuoC, NuoD, and NuoI constitute the connecting fragment. The mitochondrial homologs NuoA, NuoH, NuoJ, NuoK, NuoL, NuoM, and NuoN constitute the membrane fragment (29). E. coli complex I likely evolved by fusion of preexisting protein assemblies constituting modules for electron transfer and proton translocation (17–19, 30). Open in a separate windowFIG. 1Schematic of E. coli complex I and the nuo locus. Adapted with permission of the publisher (17, 29, 30, 49). (A) E. coli complex I is comprised of three distinct fragments: the peripheral (light gray), connecting (white), and membrane (dark gray) fragments (17, 29). The peripheral fragment (NDF) is comprised of the nuclear homologs NuoE, -F, and -G and exhibits NADH dehydrogenase activity that oxidizes NADH to NAD⁺; the connecting fragment is comprised of the nuclear homologs NuoB, -C, -D, and -I; and the membrane fragment is comprised of the mitochondrial homologs NuoA, -H, and -J to -N and catalyzes ubiquinone (Q) to its reduced form (QH₂). FMN, flavin mononucleotide. (B) The E. coli nuo locus encodes the 14 Nuo subunits that constitute complex I. The 5′ half of the locus contains a previously identified promoter (nuoP) and the majority of genes that encode the peripheral and connecting subunits (light gray and white, respectively). The 3′ half of the locus contains the majority of the genes encoding the membrane subunits (dark gray). The 3′ end of nuoG encodes a C-Terminal region (CTR) of the NuoG subunit (hatched).Because of its smaller size and relative simplicity, researchers recently have begun to utilize complex I of E. coli, and that of its close relative S. typhimurium, to identify and characterize the mechanism(s) by which cells regulate the synthesis and assembly of this large respiratory complex (3, 8, 46) and to investigate the diverse physiological consequences caused by defects in this enzyme (4, 6, 10, 40, 59). Such defects affect the ability of cells to perform chemotaxis (40), to grow on certain carbon sources (4, 6, 10, 40, 57), to survive stationary phase (59), to perform energy-dependent proteolysis (4), to regulate the expression of at least one gene (32), and to maintain virulence (5).To begin dissecting the processes by which E. coli cells regulate the expression of nuo and the assembly of complex I, we undertook a genetic analysis of the nuo locus. Here, we present the results of studies, performed on an isogenic collection of nuo mutants, that focus on the physiological, biochemical, and molecular consequences caused by the lack of or defects in several Nuo subunits. In particular, we present evidence that NuoG, a peripheral subunit, is essential for complex I function and that it plays a role in the regulation of nuo expression and/or the assembly of complex I.     

A Simple and Effective Cleavable Linker for Chemical Proteomics Applications

The study of metabolically labeled or probe-modified proteins is an important area in chemical proteomics. Isolation and purification of the protein targets is a necessary step before MS identification. The biotin-streptavidin system is widely used in this process, but the harsh denaturing conditions also release natively biotinylated proteins and non-selectively bound proteins. A cleavable linker strategy is a promising approach for solving this problem. Though several cleavable linkers have been developed and tested, an efficient, easily synthesized, and inexpensive cleavable linker is a desirable addition to the proteomics toolbox. Here, we describe the chemical proteomics application of a vicinal diol cleavable linker. Through easy-to-handle chemistry we incorporate this linker into an activity-based probe and a biotin alkyne tag amenable for bioorthogonal ligation. With these reagents, background protein identifications are significantly reduced relative to standard on-bead digestion.The covalent modification of proteins by small molecules within a complex proteome is a major theme in chemical biology and proteomics. An effective method for the detection of posttranslational modifications of proteins is the metabolic incorporation of modified biomolecules such as tagged carbohydrates or lipids (1). Reversible interactions of enzyme inhibitors, natural products, or drugs can be detected when one appends photocrosslinking agents, thereby facilitating target discovery (2, 3). A particularly interesting example of protein labeling is activity-based protein profiling (ABPP)¹ (4, 5), which utilizes the intrinsic catalytic activity of a target enzyme for the covalent attachment of an affinity or visualization tag. ABPP makes use of small molecules (activity-based probes (ABPs)) that react with the active form of a specific enzyme or enzyme class by means of a “warhead,” which is often derived from a mechanism-based enzyme inhibitor (Fig. 1A). DCG-04, for example, is based on the naturally occurring inhibitor E-64 and targets the papain family of cysteine proteases via covalent attachment of the epoxysuccinate group to the active site cysteine (Fig. 1B) (6).Open in a separate windowFig. 1.The cleavable linker strategy in ABPP. A, the elements of an ABP. B, the example ABP DCG-04, an epoxysuccinate-containing probe for clan CA cysteine proteases. DCG-04 is based on the naturally occurring protease inhibitor E-64. C, schematic strategy of cleavable linker-mediated target identification. D, the cleavage mechanism of a vicinal diol.Bulky fluorophore or biotin tags on chemical probes might interfere with efficient protein binding. Moreover, they can negatively influence the cell permeability of probes, which therefore limits their applicability in in vitro experiments. Bioorthogonal chemistries, such as the Bertozzi-Staudinger ligation (7) and the 1,3-bipolar cycloaddition of an azide and an alkyne (click chemistry) (8), allow tandem labeling strategies in which a biotin or a fluorophore is attached to an enzyme probe complex in a separate step. Consequently, the probes themselves only carry azide or alkyne groups as “mini-tags.” Tandem labeling using bioorthogonal chemistry has now become a widely used strategy to label biomolecules in lysates and in live cells (9–11).An essential step in ABPP, as well as in other chemical proteomics approaches, is the elucidation of the tagged proteins. This usually involves a biotin-mediated enrichment step followed by mass-spectrometry-based identification. Although the streptavidin-biotin interaction allows efficient enrichment as a result of the strong binding affinity (K_d ∼ 10⁻¹⁵ m), it also has limitations. The quantitative elution of biotinylated proteins requires harsh conditions (12), which lead to contamination of the sample by endogenous biotinylated and non-specifically bound proteins. These other proteins will be identified together with the real protein targets. Given that subsequent target validation with secondary assays can be a costly and time-consuming process, a reduction in false positive identifications is highly desirable. For cleaner protein identification, cleavable linker strategies (13) that allow the selective release of target proteins have been developed (Fig. 1C). The commercially available disulfide linker can be cleaved under mild conditions, but it suffers from premature cleavage in reducing media such as the intracellular environment and reducing buffers used for click chemistry and in vitro reactions of cysteine proteases. Therefore, a variety of alternative linkers for proteomics applications have been reported, including a sterically hindered disulfide (14), diazobenzenes (15–19), hydrazones (20, 21), silanes (22), light sensitive linkers (23–25), tobacco etch virus protease sensitive linkers (26, 27), and a levulinoyl-based linker (28). The synthesis of some of these linkers is lengthy or difficult to scale up, which limits their general application in chemical proteomics.Ideally, a cleavable linker is stable under a wide variety of conditions, is efficiently and selectively cleaved, and can be synthesized in a low number of easy chemical transformations. We aimed to meet these requirements by using a vicinal diol as a cleavable linker system. When vicinal diols are treated with sodium periodate (NaIO₄), the carbon–carbon bond is cleaved (Fig. 1D). Periodate treatment of proteins can result in side-reactions, such as the cleavage of linked carbohydrates or the oxidation of N-terminal serine and threonine residues. However, these N-termini rarely occur in proteins and are therefore of minor concern. In general, the mild, neutral conditions of periodate cleavage are compatible with proteins. This has been illustrated in the past, for example, by its application in the detection of protein–protein interactions (29) and the creation of unliganded MHC class I molecules (30). In this article, we report the chemical proteomics application of diol cleavable linker probes. We show that the synthesis of the linker and its probe derivatives is straightforward, that the linker is compatible with tandem click labeling, that enrichment and release of probe targets is efficient, and that the identification of targets takes place with significantly lower background than in on-bead digestion protocols.     

Activation of Heterologous Gene Expression by the Large Isoform of Hepatitis Delta Antigen

Hepatitis delta virus (HDV) encodes two isoforms of its principal gene product, hepatitis delta antigen (HDAg). These two forms play distinctive and complementary roles in viral replication. Here we report that the large (LHDAg), but not the small (SHDAg), isoform of HDAg has the capacity to activate the expression of cotransfected genes driven by a variety of promoters, including the pre-S, S, and C promoters of hepatitis B virus. Mutational analysis of the C-terminal 19 amino acids unique to LHDAg shows that changing prolines to alanines in the two PXXP motifs in this region specifically ablates the activation function without abolishing another activity of LHDAg, namely, its ability to inhibit HDV RNA synthesis. However, C-terminal truncations that also disrupt these PXXP motifs only slightly diminished the activation function, indicating that the proline mutations were not acting by inactivating potential SH3 interactions that could be mediated by these motifs. Mutation of the isoprenylated cysteine to serine decreases but does not abolish the activation activity, and overexpression of SHDAg does not interfere with the transactivation function of LHDAg. Although the mechanism and biological significance of this activity of LHDAg remain unknown, the presence of this activity serves as yet another marker that functionally distinguishes this protein from the closely related isoform SHDAg.Hepatitis delta virus (HDV) is an RNA virus that requires coinfection with hepatitis B virus (HBV) to complete its life cycle. The helper function supplied by HBV is limited to the provision of envelope proteins (hepatitis B surface antigens) for the completion of HDV assembly (28, 29, 31). HDV RNA replication is independent of its HBV helper (19). In fact, the presence of HDV suppresses HBV replication in vivo (30, 39). Nonetheless, clinical studies have shown that HDV infection can be associated with more severe hepatitis than HBV alone and is often implicated in cases of fulminant hepatitis (4, 32).The genome of HDV is a circular, single-stranded RNA of about 1,700 nucleotides (nt), of which approximately 70% are self-complementary (for a review, see references 20 and 21). This self-complemetarity allows the genome to form an unbranched rod-like structure. A unique functional protein, hepatitis delta antigen (HDAg), is encoded by the genome (3, 38), and two isoforms of this protein are produced during infection. The canonical small form of HDAg (SHDAg) is 195 amino acids (aa) long; it harbors an N-terminal coiled-coil domain responsible for oligomerization (37), a central domain responsible for binding to the RNA genome (7, 23), a nuclear localization signal (2, 7), and a C-terminal glycine- and proline-rich region with an uncertain function. This form of HDAg is essential for viral RNA replication, although it is not itself a polymerase. Host RNA polymerase II is thought to supply the polymerase function for replication (15, 26). During viral replication, an RNA editing event occurs at the UAG termination codon of SHDAg, allowing readthrough of another 19 aa (Fig. ​(Fig.1)1) to generate the large isoform of the protein, LHDAg (25). Since LHDAg contains all of the domains of SHDAg, it too can form multimers with itself and with the SHDAg isoform, bind HDV RNA (as a homo- or heteromultimer), and be localized to the nucleus. Open in a separate windowFIG. 1Sequence of the 19 aa unique to the C terminus of LHDAg. The PXXP motifs are underlined. Below are shown the amino acid changes present in the mutants employed in this study. The positions of the termination codons introduced into the truncation mutants are indicated by asterisks.Despite these similarities, the two HDAgs have very distinct functions (22) and play complementary roles in HDV replication, which takes place largely in the nuclei of infected cells (34). While SHDAg activates HDV RNA replication, LHDAg is a trans-dominant inhibitor of this process (8). By contrast, LHDAg, but not SHDAg, is capable of interacting with the HBV envelope proteins to mediate envelopment of the HDV ribonucleoprotein in viral assembly (6). This interaction has been shown to require farnesylation of a cysteine residue found in the C-terminal 19 aa unique to LHDAg (27, 16). Furthermore, it has been shown recently that only LHDAg is phosphorylated in cells (1).In this report, we describe yet another activity of LHDAg that further differentiates it from the related isoform SHDAg, i.e., the ability to activate gene expression in trans.     

Less is More: Membrane Protein Digestion Beyond Urea–Trypsin Solution for Next-level Proteomics

The goal of next-level bottom-up membrane proteomics is protein function investigation, via high-coverage high-throughput peptide-centric quantitation of expression, modifications and dynamic structures at systems scale. Yet efficient digestion of mammalian membrane proteins presents a daunting barrier, and prevalent day-long urea–trypsin in-solution digestion proved insufficient to reach this goal. Many efforts contributed incremental advances over past years, but involved protein denaturation that disconnected measurement from functional states. Beyond denaturation, the recent discovery of structure/proteomics omni-compatible detergent n-dodecyl-β-d-maltopyranoside, combined with pepsin and PNGase F columns, enabled breakthroughs in membrane protein digestion: a 2010 DDM-low-TCEP (DLT) method for H/D-exchange (HDX) using human G protein-coupled receptor, and a 2015 flow/detergent-facilitated protease and de-PTM digestions (FDD) for integrative deep sequencing and quantitation using full-length human ion channel complex. Distinguishing protein solubilization from denaturation, protease digestion reliability from theoretical specificity, and reduction from alkylation, these methods shifted day(s)-long paradigms into minutes, and afforded fully automatable (HDX)-protein-peptide-(tandem mass tag)-HPLC pipelines to instantly measure functional proteins at deep coverage, high peptide reproducibility, low artifacts and minimal leakage. Promoting—not destroying—structures and activities harnessed membrane proteins for the next-level streamlined functional proteomics. This review analyzes recent advances in membrane protein digestion methods and highlights critical discoveries for future proteomics.The goal of proteomics has grown out of matured protein identification toward the next level of function discovery, particularly including membrane proteins, through quantitative and structural proteomics at high coverage, high throughput and systems scale. Bottom-up membrane proteomics provides two paths to spearhead toward this goal. Path 1, to identify and quantify, at higher accuracy, proteins present in a mixture based on sampling fragments of each (modification-bearing or not), often aiming to find protein candidates for new biomarkers or drug targets. Path 1 represents the predominant pursuit in current membrane and global proteomics research and reviews. Path 2, to define, at atomic precision, the complete molecular identities of signaling-pivot membrane proteins—that are established high-priority therapeutic targets—under functional states and to comprehensively quantify their dynamic changes upon stimuli (Fig. 1). Path 2 includes concrete quantitation of pan-post-translational modification (PTM)¹ percent site occupancy and structural labels such as H/D-exchange (HDX), both requiring reproducible peptides that cover near-full protein sequence. This approach complements crystallography, electron microscopy (EM), NMR, and top-down MS, can discover critical details and landscapes beyond current reach, yet is largely untapped—for technical hurdles (1, 2), thus this review focuses on Path 2. As downstream state-of-the-art HPLC and mass spectrometry technologies grow mature, generic, and widely accessible, digestion sample preparation methods for membrane proteins increasingly delimit the capacity of bottom-up proteomics.Open in a separate windowFig. 1.Bottom-up quantitative membrane proteomics may serve as a hub that connects various structure and function technologies, to accelerate discovery of the structure–function mechanisms of signaling TM proteins for better therapeutics (Path 2). Breaking the barriers against direct proteomic analysis of functional-state membrane proteins, at high peptide reproducibility and coverage, is key to reaching this goal. Brackets indicate optional but often preferred steps.Abundant in human genome and pivotal in cell signaling, transmembrane (TM) proteins are coveted therapeutic targets (3), yet tremendously difficult to study at all levels (1)—including proteomics (2, 4, 5). Nearly 25% of 29,375 unique protein sequences in human proteome contain one or more TM helices (3). Prominently, TM neurotransmitter receptors such as G protein-coupled receptors (GPCR) and ligand-gated ion channels (LGIC)—with most members yet to tap—exceed 50% of current therapeutic targets (3). However, hydrophobic and prone to aggregation, membrane proteins have long vexed bottom-up proteomics with under-representation in every metric, including: sequence coverage, peptide spectrum matches (PSMs), unique peptides, and peptide reproducibility (2, 4, 5)—regardless of global lysates or highly purified samples—contrasting facile soluble proteins.Aggregation, though originated from hydrophobic TM domains, engages entire proteins and diminishes overall protease access, thus the chances to capture non-TM peptides at authentic quantity from aggregated samples are proven slim and unpredictable, and this fact cannot be reversed by simply changing database size. Global analysis without the fair share of membrane proteins is hardly global, most readily detectable soluble proteins do not make useful new drug targets, and sporadic sketchy touches on TM receptors, known high-priority targets, generate no concrete structure–function roadmap to guide therapeutic discovery. Therefore, for the next-level functional proteomics aiming to improve health, the technical barriers of membrane proteomics must be solved.Most digestion methods emphasized finding more proteins from mixtures, yet few addressed comprehensive peptide reproducibility, let alone both. Although identifying the same membrane proteins—represented by two or more unique peptides—between independent digestions and sample states was routinely achieved in shotgun proteomics, until recently (6, 7), comprehensively reproducing peptide forms and abundances, which matters for peptide-centric peak-area or tandem mass tag (TMT)-based quantitation of expression, modification, and labels, remained out of reach (2, 4, 5).The challenges in detecting peptides for membrane proteins are twofold: (1) forming peptides, and (2) detecting formed peptides. Using prevalent overnight urea–trypsin in-solution digestion, extreme three-dimensional 8-day HPLC separations benchmarked sequence coverage at 20–30% for membrane proteins (8), demonstrating deficient peptide formation pre-HPLC is a major unmet need, although many recent efforts to delve deeper focused on HPLC, mass spectrometer, and tandem MS strategies downstream to digestion (8, 9). Further, unaddressed PTMs of mammalian membrane proteins (such as extensive glycosylation at high site occupancy (10)), may prevent sequence identification, even if peptides are formed. Both hydrophobic TM domains and intricate PTMs, hallmarks of human signaling TM proteins, are important for functions. Detergents, inherited from protein extraction (typically SDS and Triton X-100), severely challenged downstream digestion and/or HPLC steps in global proteomics.The “urea–trypsin solution” tradition contains three elements—urea denaturation, trypsin, and in-solution format—that each and all, deserve critical thinking for membrane proteins. Myriad resorts have been proposed to address membrane protein gel-free digestions for various needs over past decades (11, 12), yet attempts to denature and subdue proved ineffective. This review will focus on advances since 2009 for prospects to support the next-level streamlined high-coverage, high-throughput quantitative membrane proteomics (Open in a separate windowAdvances of membrane protein gel-free digestion methods. MSP, membrane scaffold protein; FDD, flow/detergent-facilitated protease and de-PTM digestions for deep sequencing. st, StageTip; iodoaa, iodoacetamide; ovnt, overnight (14 h); N-glyco, N-glycosylation; med, medium; id, identify; qt, quantify; hp, human proteome search^a Sequence coverage also depends on HPLC method and sequence database used for search.^b Purified protein searched against specified sequence.^c Identified by searching against human proteome (original, downloaded May, 23, 2013 from Ensembl) at peptide FDR <1%, using SEQUEST-HT and Percolator in PD 1.4. Signaling peptide sequence was not removed (a “/95%” correction, assuming a 20-residue signaling peptide for a total 500-residue protein). MSP, membrane scaffold protein; FDD, flow/detergent-facilitated protease and de-PTM digestions for deep sequencing; st, StageTip; iodoaa, iodoacetamide; ovnt, overnight (14 h); N-glyco, N-glycosylation; med, medium; id, identify; qt, quantify; hp, human proteome search.

In-solution Digestion and its Limits

Although prevalent, urea–trypsin in-solution digestion is neither complete nor predictable (13) for membrane proteins, and poses a weak link to future proteomics. Typical workflow includes: protein extraction by boiling 4–7% SDS, acetone or acid precipitation to remove SDS, resolubilization in 8 m urea, DTT reduction, iodoacetamide alkylation, in-solution digestion with trypsin in 2 m urea overnight, peptide desalting with C18 to remove the high urea, peptide drying to remove C18 elution solvent and concentrate (often causing oxidation), and redissolving in HPLC sample buffer. Pronounced problems are: serious unpredictable protein loss to aggregation (14), high oxidation artifacts and contaminations introduced by laborious processing (15), biases over protease specificity and abundance (13, 16), low digestion efficiency, coverage, and peptide reproducibility (2, 4).Previous efforts to improve membrane protein digestion have focused on proteases with strict specificity, such as trypsin, Lys-C, Glu-C, Glu-N, and Asp-N, aiming to concentrate the quantity of peptides in defined theoretical forms. However, applied with denaturation and/or protease-deficient in-solution conditions, the advantage of this strategy was masked by large loss of accessible membrane proteins to aggregation (including re-aggregation), as evidenced by previous sub-25% coverage of purified human GPCR β₂-adrenergic receptor (β₂AR) using specific or nonspecific proteases in urea (6), and by common low numbers of peptides identified for Cys-loop LGICs from multiple rat or mouse brains using high-urea Lys-C and/or trypsin in-solution digestions. Further, in-solution digestion requires low protease-to-substrate ratio (1:50 or lower, m/m) to reduce contamination, thereby operates near-entirely in the enzyme-deficient, diffusion-limited, low-efficiency region (7); slow reaction rates of compromised trypsin in urea amplify such deviation from ideal. In-solution trypsin (2 m urea) digestion has also been extended by combining with other enzymes such as Lys-C (8 m urea) (17), pepsin (18), Asp-N (19), and PNGase F (pre-digestion (19)). Increasing evidence suggests multispecificity, often via using multiple proteases, is necessary for reliable PSM quantitation of lysates (13, 20). Pressure cycling technology speeded tandem urea-solution Lys-C/trypsin digestion of tissue lysate into 6 h, and strict cycle control enhanced peptide reproducibility (21).

Alternatives to Traditional In-solution Digestion

On-filter In-solution Digestions (FASP and in-StageTip)

Avoiding complete precipitation/resolubilization, the filter-based in-solution digestion method FASP (22, 23) achieved landmark improvement in digestion yield (up to 50%) by keeping membrane proteins in solutions during SDS removal. Following SDS protein solubilization, FASP applied dialysis (dial)-filtration with small molecular-weight-cut-off filters to gradually replace SDS in solution with high urea, performed in-solution digestion(s) overnight on filter, then separated peptides from enzyme and undigested proteins via centrifugation. FASP and its derivatives that added deoxycholic acid (2014 eFASP (24)) or further extended the on-filter reactor for reactions such as deglycosylation (2010 N-glyco-FASP (25)), have been widely applied since 2009. However, ∼50% protein sample resists digestion likely because of inevitable re-aggregation upon detergent depletion, and although overall protein identification is improved, peptide reproducibility remains low (23). The laborious manual operation is prone to contamination and sample loss. Abandoning SDS, the latest version of FASP, 2014 in-StageTip (26), used smaller tips sealed with C18 disk that doubles as micro-filter besides desalting, and combined protein deoxycholate solubilization, reduction, alkylation, and overnight in-solution digestion in these tips in 96-well format. However, this method still mandates manual processing, hours of digestion, and centrifugation (which interrupt automation streamline), and inherits all problems intrinsic to the denaturation-trypsin in-solution paradigm; ionic and high in cmc (critical micelle concentration; 1% m/v, 4.2× cmc was 25 mm), sodium deoxycholate incurred an extra step of post-digestion removal with ethyl acetate extraction and centrifugation (26), further obstructing automation.

Protein On-bead Digestions

Fixing membrane proteins on beads following specific affinity-enrichment (27, 28) effectively alleviated interprotein aggregation, such as during detergent removal (27), and greatly improved digestion and identification of membrane proteins using Lys-C and trypsin urea solutions (27). However, information of unbound proteins is lost. In the 2014 Single-Pot Solid-Phase-Enhanced Sample Preparation (SP3) method, hydrophilic magnetic beads adsorbed proteins and peptides less discriminatingly, and simplified solution separation for pre-digestion SDS removal and post-digestion peptide cleanup (29). However, common to both protein on-bead strategies, inevitable intraprotein aggregation may limit digestion access, and peptides remain mixed with protease solution.

Immobilized Enzyme Reactors for Fast Reproducible Digestion

Numerous micro-reactors of enzymes, including trypsin (30–32), trypsin/Lys-C (33), pepsin (34, 35), and PNGase F (36–38), adsorbed or covalent-bonded on various solid supports, have been described for peptides and/or glycans studies, mainly for purposes of faster reaction and easy product separation. They demonstrated advantages over in-solution methods, but mostly using simple soluble proteins in high denaturants such as urea, guanidine, or organic solvent.Covalently immobilized pepsin columns emerged in 2002 for HDX to rapidly digest proteins at low pH low temperature to minimize D-label back-exchange, without complicating spectra with protease peaks (34). With high urea or guanidine denaturant (1.5–3 M, some studies also added 500 mm TCEP) at pH 2.5, pepsin column successfully digested most soluble proteins tested at high (over 90%) coverage and high peptide reproducibility within seconds-minutes, and is widely adopted in HDX (34, 35, 39, 40). HPLC-grade pepsin column''s advantages are multifold and attributable to its operation as a high-surface-concentration plug-flow reactor, in retrospect (7). First, immobilizing pepsin on POROS beads can increase effective surface concentration to over-1-mm scale (34), near-thousand-fold higher than the 2 μm (or 0.1 mg/ml, 50 kDa) protein typically applied, thereby allowing pepsin column to operate well within the abundance-unbiased high-efficiency region, unrestricted by total sample size (7). Second, flow drive overcomes the problems of product inhibition caused by slow diffusion intrinsic to in-solution incubation, and affords a precise control of the fast product formation from nonspecific protease, which is unconceivable to in-solution and beads-based methods. Third, typical column size, flow rate (1–2 mm i.d. × 20 mm, 25–200 μl/min (6)) and pressure fit with HPLC system for full automation. Trypsin POROS column also showed efficient real-time digestion of soluble proteins online in organic solvent (30–32). However, until 2010 (6), protease columns met no success with membrane proteins, and for years GPCR β₂AR coverage remained below 25% (6, 41), preventing useful HDX mapping.

Alternative Detergents for Protein Extraction and Digestion

Other advances were achieved by changing detergents. Traditional detergents for protein extraction in proteomics, such as ionic SDS and poly-disperse Triton X-100, inhibit protease digestion and/or peptide HPLC MS, but are difficult to remove. Alternatively, novel protease-compatible use-and-shred detergents such as RapiGest (42), could increase membrane protein sequence coverage from 10–20% to about 30% in global proteomics (42), and are increasingly applied with success in place of, or after (hours of dial-filtration), SDS or Triton extraction since 2007. However, these detergents vary solubilization power with proteins (some instantly precipitate proteins such as cytochrome c oxidase, CcO), require chemical cleavage post-digestion—difficult for fast full automation, and some cleaved products contain amines that interfere with TMT. A latest acid-cleavable analog of SDS, anionic MaSDeS, showed SDS-like strong solubilization power on tissues, though it is severely denaturing and forms amine products (43, 44). Amphi-polymers (amphipols) can support protein extraction and digestion, but most aggregate below pH 3–4 and require removal pre-HPLC by precipitation and centrifugation (45), interrupting automatable workflow. Several recent studies compared numerous detergents for digestion, but most remained confined to industrial poly-disperse and ionic detergents that deactivated proteins and required removal; other attempts combined harsh detergents, organic solvents, high urea, high pressure, and high temperature (46).Despite these incremental advances in membrane protein coverage, obvious obstacles endured. First, hydrophobicity and aggregation persisted. Second, PTMs remained mostly unaddressed. Third, all these methods incur laborious manual operation, centrifugation (except for magnetic beads) and hours of processing before HPLC injection, interrupt automation, and remain unable to meet the minutes'' time window allowed by HDX pipeline. Central to these method designs are the common observation and belief that: Detergents are incompatible with the protein-peptide-RP HPLC-ESI MS/MS pipeline, and must be avoided or removed at some point pre-HPLC. Further, the concepts of protein solubilization versus denaturation, and protease digestion reliability versus theoretical specificity, were frankly undistinguished. Consequently, Path 2 (Fig. 1), direct pan-PTM and HDX structural mapping of native functional human membrane proteins—the established high-priority therapeutic targets urgently awaiting precise structure-function roadmaps—remained a formidable field to bottom-up proteomics.

Detergent Selection for Functional Proteomics: The Schrödinger''s Cat Scare

For protein states, we see what the method presents. As a primary tool to retrieve TM proteins from lipid bilayers for solution-based analysis, detergents present proteins in detergent-protein micelles, and inevitably—more or less—affect their function and structure integrity. Rather than presolubilization cross-linking that may disturb the system and complicate analysis, functional proteomics predominantly counts on detergents to preserve proteins'' authentic conformation and non-covalent interacting network, during solubilization and enrichment. Thus ideal detergents ought to: (1) solubilize membrane proteins, (2) maintain proteins'' native structure and interactome, and (3) reduce downstream cleanup that loses samples to re-aggregation and biases against hydrophobic components.Although current proteomics literature frequently mixed detergents'' solubilization strength with denaturation severity (47), they are in fact two distinct concepts—a premise for functional proteomics. Detergents vary vastly in their effects on protein activity despite solubilization (4, 5, 22), at unavoidable sample loss.

Table II

Open in a separate windowDetergent selection for functional membrane proteomics. cmc, critical micelle concentration; AN, aggregation number; dtg, detergent; n/a, not available. DM, n-decyl-β-d-maltopyranoside; OG, n-octyl-β-d-glucoside. FA, steroid-based facial amphiphiles; FA-3, type 3 (2 maltosides); FA-4, type 4 (3 maltosides); FA-5, type 5 (3 glucosides). BP-1, beta-strand peptide type 1, (N) acetyl-(octyl)G-S-l-S-(N-methyl)l-d-(octyl)G-d-NH₂^a Source for cmc and aggregation number is Anatrace (A), Thermo Pierce (T), or specified reference.^b Tissue-, cell- and protein-dependent, shown are general trends, based on Refs (6, 7, 24, 45, 49, 51–53, 61, 66–70), detergent charge state, count ratio over protein, PDB database, and experiences with CcO, hGPCR β₂AR and hLGIC GABA_AR. Utility/Compatibility: Extraction from membrane (E; success rate E > e; e: requiring sonication, or effective for mammalian and insect cells but less effective for bacterial cells), Keeping protein soluble (K), Activity maintenance (A; AA > Aa > A > a). Crystallography (X; X > x), EM (EM > em; em: often difficult to tell protein TM domains from detergent micelles), Digestion (D; D > d), and peptide RP HPLC MS (M; M > m). Unmarked means no or usually unconsidered.^c After exhaustively replacing other detergents.cmc, critical micelle concentration; AN, aggregation number; dtg, detergent; n/a, not available; DM, n-decyl-β-D-maltopyranoside; OG, n-octyl-β-D-glucoside; FA, steroid-based facial amphiphiles; FA-3, type 3 (2 maltosides); FA-4, type 4 (3 maltosides); FA-5, type 5 (3 glucosides); BP-1, beta-strand peptide type 1, (N) acetyl-(octyl)G-S-L-S-(N-methyl)L-D-(octyl)G-D-NH2.Rigorous membrane protein functional interactome studies typically compared multiple detergents—as they produced vastly variable, partially overlapping protein populations—and emphasized proteins that overlapped or were unique (48, 49). Subsequent analyses resorted to detergent removal by overnight RapiGest exchange-cleavage (48) or by protein precipitation-resolubilization (49), and digestion with trypsin solution (48) or beads (49).Broadly defined as “a reagent to wipe away” and chemically also termed surface-active agent (surfactant), detergent is characterized with being amphiphilic, and the hydrophilic domain can assume various shapes: one pole of a rod (SDS, Triton X-100), one face of a bulky molecule (CHAPS, deoxycholate), or one side of a linear molecule (amphipols) (d-maltopyranoside (DDM) was synthesized and first shown to be unusually effective for both solubilizing and stabilizing active membrane proteins—attributable to its micelle size and rigid tight-packing nonionic head group—by the Ferguson–Miller group in the 1980s (50–53), using animal tissues then cell cultures. Subsequently, DDM-based extraction and/or purification have critically enabled high-resolution crystallization of most of the over-500 diverse membrane proteins in the Protein Data Bank, including GPCRs, multisubunit ion channels, neurotransmitter transporters, and respiratory complexes (54–60); DDM''s recent “X”-shaped dimeric derivative, maltose-neopentyl glycols (MNG) type 3 (MNG-3, lauryl MNG), stabilized hGPCR-Gs complex for crystallization (55, 61). For mammalian signaling TM proteins native to ∼20–25%-cholesterol lipid bilayer habitats (by weight and by count) (62–64), the Stevens group found adding cholesteryl hemisuccinate (CHS) to DDM forms wider bicelle-like DDM/CHS micelles that crucially stabilize human GPCRs for activity and crystallization (54, 65). DDM/CHS also proved effective to extract and purify active synaptic hetero-pentamer hGABA_AR from HEK293 (66); seconds of DDM/CHS sonication afforded over 90% solubilization of HEK293 membranes (66) and no visible pellet post-centrifuge for rat brains (personal communication, Feb 2015). Decyl MNG/CHS stabilized hLGIC (β3)₅ GABA_AR for crystallization (67). By contrast, traditional ionic or poly-disperse detergents, such as SDS, CHAPS, cholate, deoxycholate, C12E9, Triton X-100, Tween 20, and Brij 35, as well as nonionic DM, OG, and digitonin, failed to emulate despite decades of pursuit (51–53, 68). Ionic CHAPS and cholate consistently deactivated hGABA_AR (66) and mammalian CcO (52) by several-fold compared with DDM and DDM/CHS. Latest nonionic detergents that combined two to three DDM''s head groups or three glucosides with a steroid body markedly stabilized proteins for crystallization (68). By replacing other detergents, various amphipols and beta-strand peptides also emerged effective to chaperone proteins for high-resolution EM (69, 70).However, these detergent-driven advances in membrane protein technology since 1980 circulated largely among the crystallography community, not proteomics. Inheriting industrial-grade detergents such as SDS and Triton X-100, historically applied for cheap price, proteomics has suffered huge penalties in quality and cost. Using ESI MS experiments of intact model protein solutions (infusion and flow injection), a pioneering 1994 study reported several nonionic, zwitterionic, and anionic detergents tolerable by ESI MS, in acetic acid/acetonitrile/H₂O spray (top 7: n-dodecyl glucoside, n-hexyl glucoside, CHAPS, cholate, and three equally scored DDM, OG, and octyl thioglucoside), and in H₂O spray (top 5: n-dodecyl glucoside, n-hexyl glucoside, OG, n-octyl sucrose, and n-dodecyl sucrose) (71).We further considered detergent compatibility with protein extraction, protein activity, protease activity, RP-HPLC, data-dependent MS/MS competition and peptide labeling. We propose that beyond tolerance by intact protein MS, detergents may be cultivated as a tool for both membrane protein extraction/purification and digestion modules in proteomic pipeline. Detergents as a tool for membrane protein extraction and purification (discussed above) emphasize maintaining protein native conformations and activity at high yield (often with additives), and providing near-equal grounds of protein states to correlate with other technologies such as crystallography, EM, and function assays (Fig. 1). Detergents as a tool for digestion put emphasis on maintaining protease activity, solubilizing and stabilizing substrates against inter- and intraprotein aggregation over a wide pH range, and minimizing interference with labels (such as HDX, oxidation and TMT), peptide RP HPLC, or ESI MS/MS, but less on native conformations, as digestion often desires acidic pH and proteins are dissected rapidly. Detergents as a tool for united proteomic pipelines emphasize optimal compatibility with all modules.Based on detergent properties—including charge state at acidic pH, mono- or poly-dispersion, and cmc (count ratio over protein at 5× cmc:1 μm protein)—and their documented performances in membrane protein extraction, activity maintenance, and high-resolution structures, particularly experiences gained from CcO, hGPCR, and hLGIC complexes, Direct Flow/DDM-facilitated Digestions for HDX, Deep Sequencing, and Quantitation

Discovery of Omnicompatible Detergents as a Pivotal Tool to Fully Automate Deep and Direct Membrane Proteomics

Breakthroughs in membrane protein digestion started in 2010 for HDX dynamic structure studies, arguably the ultimate digestion challenge, by including detergents that complied with structure/protease/HPLC MS (6), or by applying membrane scaffold proteins (MSP) that kept proteins in nanodiscs followed by cholate (which entailed high-pressure ultra-performance liquid chromatography, UPLC) (72, 73), to solve aggregation. Thinking outside the box of denaturation anddetergent removal, we discovered that nonionic low-cmc amine-free DDM—the most protein nature-promoting and crystallization-successful detergent to date—was completely compatible with structure/protease/TMT/HPLC MS/MS, and devised DDM-based protein extraction, purification and digestion methods that cleared all these challenges of membrane proteins (6, 7, 66). DDM-low-TCEP (DLT, for HDX) and flow/detergent-facilitated protease and de-PTM digestions (FDD, for deep sequencing and quantitation) both completed peptide preparations within seconds-minutes, at robust coverage and peptide reproducibility supporting full automation, demonstrated by using hGPCR (6) and hLGIC complex (7), respectively.Advantages of DDM as a tool for membrane proteomics are multifold and pivotal to achieving fully automated deep and direct proteomic analyses of membrane proteins in physiological states. First, for protein extraction/purification, DDM combined with CHS to mimic cholesterol in human cells, can extract membrane proteins at high activity, and high efficiency when applied with co-micelle-promoting slow addition or sonication (66). DDM and its derivatives that critically enabled the purification and crystallization of myriad active membrane proteins (50, 51, 54–57, 61, 74, 75), present proteins in functional states in solution, thus providing a common ground for proteomics to bridge functions directly with protein PTMs, dynamic conformations, and static structures resolved by crystallography (Fig. 1). Second, as a tool for digestion, DDM preserves—not destroys—the activity and solubility of both enzymes and protein substrates, thus facilitates effective catalytic contact and unbiased flow of hydrophobic species during digestion, and obviates problematic re-aggregation, high urea, subsequent desalting, and reconcentrating. DDM is effective over a wide pH range, thus can broadly simplify diverse digestion formats such as: enzyme column, protein on-bead, in-solution, and in-nanodisc, by replacing conventional detergents, urea or guanidine. Third, for overall pipelines, amine-free nonionic DDM supports downstream HPLC, MS/MS competition and TMT reactions, can be included throughout the cell lysis-protein extraction-(HDX)-digestion-(TMT)-HPLC-MS/MS process without removal, thus confers the workflows with no interruption, no sample leakage and high sensitivity. DDM at 0.4–1 mm (0.02–0.05%, 2.3–5.7× cmc) over 1 μm proteins (0.04–0.28 mg/ml) was well tolerated (6, 7).At even lower cmc (∼9, ∼10, and 36 μm) (61), nonionic MNG-2, MNG-3, and decyl MNG are expected compatible with RP HPLC MS as well; likewise are the low-cmc nonionic facial amphiphiles (Fig. 2B and 61), we found its membrane solubilization power for HEK293 was comparable to DDM basal (66), thus MNG can serve as a valuable control to cross-check DDM extraction results and as another tool for digestion.Open in a separate windowFig. 2.General approaches of fully automatable flow-and-detergent facilitated membrane proteomics. A, Workflows of flow/detergent-facilitated protease and de-PTM digestions (FDD) and DDM-low-TCEP (DLT) for membrane protein samples, B, structures of protein activity/HDX/TMT/RP-HPLC/MS omni-compatible detergents and additives, and C, alternative modes of FDD. Instrument images were from www.thermoscientific.com.

DDM-low-TCEP Digestion for HDX

Instead of conventional 3 m urea, 1.5 m guanidine, or 500 mm TCEP, the 2010 DLT method developed a digestion solution of 0.02% (m/v) DDM, low 15 mm TCEP and 100 mm NaH₂PO₄-HCl pH 2.4 (final 7.5 mm TCEP), to quench membrane protein HDX (DDM/CHS/glycerol protein buffer in H₂O or D₂O) for direct digestion, online in a fully automated workflow (6). DLT with pepsin column achieved the first comprehensive HDX profile of 7TM hGPCR at near 90% coverage, using a PTM-deprived minimalistic β₂AR developed for crystallization (6). These peptides were each reproduced by several hundred independent digestions of apo and various ligand-bound β₂AR from multiple batches of purifications (Ref (6) and unpublished data). The DDM solution also successfully reproduced high coverage for other GPCRs in other laboratories (76).However, devised for direct HDX, DLT did not address extensive PTMs and Cys, as they did not pose a problem in the β₂AR crystallizable construct: All PTM-bearing domains were either genetically truncated (intracellular C-terminal tail and loop 3) or deglycosylated (two adjacent extracellular-domain N-glycosylations) in purification (6). Although 7.5 mm TCEP allowed detecting peptides bearing free Cys, without alkylation, they were the weakest, and the C–C linked versions were detected at higher ion counts (6). Also designed with a desalting trap and high-flow-rate HPLC and ESI (50 μl/min), DLT-HDX''s peptide signals were close to baseline in HPLC-ion count chromatograms, and did not appeal to broader applications (Fig. 3A).Open in a separate windowFig. 3.At much higher peptide ion sensitivity, the 2015 FDD-no-trap-nESI method proved DDM can be a superior tool for broad nESI-based membrane proteomics. Representative HPLC MS traces of (A) 2010 DLT-HDX-trap-ESI of hGPCR apo β₂AR, typical to the data for (6), B, FDD-no-trap-nESI of hGABA_AR, and (C) independent digests of H/D versions of hGABA_AR using FDD-no-trap-nESI''s HDX module showing good reproducibility. All spectra were acquired with orbitrap analyzers in Thermo Exactive (A), Q-Exactive (B), or LTQ-orbitrap XL (C) mass spectrometer. Experiments used similar concentrations of protein and detergent, but A loaded several-fold more total sample to HPLC than B and C. Figure was adapted from (7).

Flow/DDM-facilitated Protease and De-PTM Digestions for Deep Sequencing and Quantitation

Because DDM obviated high salt, by removing the desalting trap and adopting capillary HPLC column and nanoESI, a no-trap-nanoESI upgrade of DLT remarkably increased peptide ion sensitivity (66). When applied to full-length hGABA_AR hallmarked with Cys-loops, it produced up to 80–84% coverage with a 75-min HPLC gradient (66), but gaps of over 15% persisted at Cys and potential PTM sites.Addressing PTMs and Cys, the 2015 FDD method further combined immobilized pepsin column, PNGase F column, Cys-selective reduction and alkylation, and greatly improved peptide metrics. A novel integrated digestion paradigm, FDD''s components each targeted the key challenges for human signaling TM proteins—hydrophobicity and intricate PTMs—inaccessible for decades by conventional methods. (1) FDD''s tandem flow reactor format achieved fast, effective, reproducible, and controllable reactions for both proteolysis and de-PTM; (2) DDM resolved aggregation caused by TM hydrophobicity and promoted unbiased flow through all reactors; (3) de-PTM PNGase F column solved thorough mapping of N-glycosylation sites that turned out to crosstalk with other PTMs and critical for functions. At minutes of total preparation time, FDD achieved up to 99% coverage in one run, high peptide reproducibility and low artifacts for all subunits of hGABA_AR/HEK293 (7) (Fig. 4A and Fig. 3C). Importantly, the FDD-no-trap-nESI platform increased peptide ion sensitivity by four orders of magnitude from 2010 DLT-trap-ESI (Fig. 3), and discovered DDM is no longer a compromise, but a superior tool broadly applicable to nESI-based proteomics. FDD further demonstrated direct TMT labeling and analysis (7) can be integrated in a fully automatable membrane proteomic workflow (Fig. 2).Open in a separate windowFig. 4.Protein sequence coverage identified from direct FDD-pepsin digestion and analysis of DDM/CHS-enriched hGABA_AR solution, by searching against. A, the target sequences of (α1)₂(β3)₂(γ2L)₁ GABA_AR (SEQUEST in Proteome Discoverer 1. 3), or (B) the original human proteome (May 23, 2013 Ensembl, SEQUEST-HT in Proteome Discoverer 1.4), both at no protease specificity and filtered to peptide FDR <1% by Percolator. Each thin line represents one PSM (apple green, α1 subunit; blue, β3; orange, γ2L); bold lines indicate domains (dark blue, TM; red, intracellular loop; black, extracellular Cys-loop). B, shows the % sequence coverage (blue bar) for the top 34 proteins (ranked by PSMs and unique peptides), and their percent PSM (red bar) of the total PSMs of all proteins identified by three or more valid unique peptides; (m) designates membrane residence, as annotated in human proteome database. N-terminal signaling peptide sequences were not removed. A was adapted from (7). These proteins were mainly cell-originated survivors of two affinity screenings by membrane isolation and by buffer washing during anti-FLAG enrichment, not contaminant from processing.FDD presents multifold advantages for bottom-up membrane proteomics, and critically supports the next-level quantitative-structural integration. First, FDD transformed the days-long aggregation-plagued paradigm into immediate digestion and measurement of active protein solutions—in seconds–minutes total peptide preparation time—at unprecedented coverage, reproducibility, and authenticity. This allowed both the minutes-windowed HDX and the deep PTM/interactome mapping to look at the near-identical states of proteins and peptides, thus broke barriers against integrating quantitative-structural membrane proteomics. Second, FDD eliminated precipitation, filtration and centrifugation central to the denaturation-, tip- or filter-based in-solution traditions. This enabled a robust, instant, versatile protein-(HDX)-peptide-(TMT)-HPLC MS streamline for uninterrupted automation. Together, FDD made Path 2 immediately achievable (Fig. 1) and may improve Path 1 in speed, depth, accuracy, sensitivity, and throughput (6), and DDM-facilitated pepsin solution, beads versus flow reactor format (66)—were compared on the equal grounds of target sequence search and parallel experiments. Second, spectra search against human proteome maintained the high peptide number and coverage of target proteins (7) (Fig. 4B). Third, identified proteins showed average sequence coverage higher than current ∼25% benchmark (8, 42) (Fig. 4B). Further, compared one-to-one with 7TM hGPCR and hGABA_AR, most proteins in cell lysates are technically easier, thus unlikely to pose further obstacles.Concrete reach to PTMs—including N-glycosylation on Cys-loop and endogenous M-oxidation—proved crucial for discovering function clues inaccessible to methods with sporadic coverage and severe artifacts (7). Extensive PTMs in human signaling TM proteins, such as high-occupancy N-glycosylation, had remained largely a target of observation, but rarely exploited as a means to aid overall peptide formation and detection, partly for fear that PTM enzymes overwhelm real samples (though PNGase F was used to release enriched glycopeptides (77)). Predigestion PNGase F solution incubation indeed identified more proteins in yeast cell wall (19). But emerging complicated N-glycosylation scenarios in mammalian TM proteins suggest, treating peptides post-digestion is more likely efficient and complete than treating proteins (7, 66).

Extended Applications of FDD Format

As a general platform to prepare membrane protein peptides, FDD''s elements—enzyme flow reactors, omnicompatible detergents, and de-PTM—are flexible to adapt to nearly all levels of proteomics and support myriad enzymes and new detergents (Fig. 2C). Both the DDM/CHS-extraction and FDD digestion strategies are compatible with upstream enrichment of cell fractions, proteins, protein nanodiscs, downstream peptide enrichment (DDM supports IMAC (6, 54, 57, 74, 75)), fractionation, TMT, and HPLC. Excitingly, besides DDM/CHS, MNGs and several recent detergents superb for crystallography and EM likely fit FDD as well (Fig. 1). This flexibility also allows matching digestion detergent with the one used in protein preparation to avoid multiple HPLC peaks.

Membrane Protein Digestion: Protease Reliability beyond Theoretical Specificity

Confounded by membrane proteins'' unique hydrophobicity and tendency to aggregate, protease''s empirical reliability has proved distinct from its theoretical specificity over primary sequence. How to apply the protease appears to matter more than theoretical specificity for achieving reliable digestion. Several recent digestion advances can be interpreted via improving substrate–protease contacts, such as the protease-mild substrate-solubilizing RapiGest (42), and the pressure cycling technology to improve reaction mixing in urea (21), though all under the detergent-removal/avoiding paradigm. For this purpose, an HPLC-grade flow-propelled high-concentration immobilized enzyme micro-column reactor is arguably the most effective. Omnicompatible detergents provided the bridge for membrane proteins to take advantage of this format, to reach complete reproducible digestion and seconds–minutes scale under ambient physiological solutions, temperature, and pressure, obviating any brutal force.For specific proteases, effective substrate-protease contacts shall help overcome the deviation from theoretical specificity, and finally achieve the “peptide concentrating” strategy described above. DDM-facilitated trypsin beads already showed complete digestion of a multihelix protein within minutes (supplemental Fig. S1A) contrasting overnight in urea solution, thus DDM-facilitated trypsin column is expected effective with membrane proteins too (Fig. 2C). Applying multiple specific proteases in the format of HPLC-grade tandem columns or mixed-beads column (minutes) shall be more efficient than combining days of individual urea-solution digestions (Fig. 2C).Challenging conventional belief, fast multi-specific protease proved able to be highly reproducible and reliable—when applied in the DDM-facilitated HPLC-grade enzyme column format (6, 7). Reproducible peptide ladders provide desired redundancy and bypass specific proteases'' cleavage gaps. Pepsin column rapidly generated comprehensive coverage including TM domains, with redundant overlapping peptides four to 20 residues long, useful for HDX and PTM mapping (6, 7). Nepenthesin-1, a recent pepsin alternative, has different multispecificities, max activity at pH 2.5, low autolysis, low tolerance to denaturants, yet stability in basic pH (78, 79)—making it suitable for high-yield immobilization and nondenaturant digestion.Fast and preferring hydrophobic sites, HPLC-grade columns of acidic multispecific proteases such as pepsin and nepenthesin-1 (78–80) offered a robust, efficient (seconds) alternative to combining multiple solutions of specific proteases (days), and can be favored over trypsin particularly: (1) when TM receptors such as hGABA_AR frequently contain K/R over 50 residues apart in extracellular domain, but also dense K/R (1–2 residues apart) in intracellular domain: Seeing these K/R-dense peptides relies on trypsin''s unreliability—missing K/R cleavage, a ∼15% chance in urea solution (81), (2) when K/R-PTMs interfere with trypsin cleavage impeding peptide-centric quantitation, (3) when proteins such as myoglobin resist trypsin digestion (supplemental Fig. S1B Lane 3), and (4) when acidic digestion is desired to minimize nonenzymatic deamidation (82) and scrambling of native PTMs such as C-C bonds (83). The large number of peptides is within capacity of matured HPLC separation.     

Endogenous 3,4-Dihydroxyphenylalanine and Dopaquinone Modifications on Protein Tyrosine: LINKS TO MITOCHONDRIALLY DERIVED OXIDATIVE STRESS VIA HYDROXYL RADICAL*

Oxidative modifications of protein tyrosines have been implicated in multiple human diseases. Among these modifications, elevations in levels of 3,4-dihydroxyphenylalanine (DOPA), a major product of hydroxyl radical addition to tyrosine, has been observed in a number of pathologies. Here we report the first proteome survey of endogenous site-specific modifications, i.e. DOPA and its further oxidation product dopaquinone in mouse brain and heart tissues. Results from LC-MS/MS analyses included 50 and 14 DOPA-modified tyrosine sites identified from brain and heart, respectively, whereas only a few nitrotyrosine-containing peptides, a more commonly studied marker of oxidative stress, were detectable, suggesting the much higher abundance for DOPA modification as compared with tyrosine nitration. Moreover, 20 and 12 dopaquinone-modified peptides were observed from brain and heart, respectively; nearly one-fourth of these peptides were also observed with DOPA modification on the same sites. For both tissues, these modifications are preferentially found in mitochondrial proteins with metal binding properties, consistent with metal-catalyzed hydroxyl radical formation from mitochondrial superoxide and hydrogen peroxide. These modifications also link to a number of mitochondrially associated and other signaling pathways. Furthermore, many of the modification sites were common sites of previously reported tyrosine phosphorylation, suggesting potential disruption of signaling pathways. Collectively, the results suggest that these modifications are linked with mitochondrially derived oxidative stress and may serve as sensitive markers for disease pathologies.Generation of reactive oxygen species (ROS)¹ and reactive nitrogen species is a normal consequence of aerobic metabolism that, in excess, results in oxidative stress that further leads to oxidative modification of proteins, lipids, and DNA, events that may lead to altered cellular function and even cell death (1, 2). Chronic oxidative stress is well recognized as having a central role in disease and is responsible for both direct alteration of biomolecular structure-function and compensatory changes in cellular processes (1–4). It is increasingly recognized that oxidative modifications of proteins can serve as potential biomarkers indicative of the physiological states and changes that occur during disease progression. Thus, the ability to quantitatively measure specific protein oxidation products has the potential to provide the means to monitor the physiological state of a tissue or organism, in particular any progression toward pathology. Given Parkinson disease (PD) as an example, a number of oxidative modifications on proteins pertinent to PD have been identified, further supporting the potential importance of oxidative modifications to disease pathogenesis (5).Many oxidative modifications on specific amino acid residues, such as protein carbonylation (6), cysteine S-nitrosylation (7–9), cysteine oxidation to sulfinic or sulfonic acid (10–12), methionine oxidation (13, 14), and tyrosine nitration (15–21) within complex protein mixtures, have been detected by MS-based proteomics; however, their low abundance levels within complex proteomes often hinder confident identification of these potentially significant modifications (22). For example, tyrosine nitration is a well studied post-translational modification mediated by peroxynitrite (ONOO⁻) or nitrogen dioxide (^·NO₂), which commonly occur in cells during oxidative stress and inflammation; however, only a small number of nitrotyrosine proteins have been identified from a given proteome sample because of insufficient analytical sensitivity and the chance of incorrect peptide assignments (19, 23). With recent advances in high resolution MS that provide high mass measurement accuracy, the ability to confidently identify modified peptides has been significantly enhanced (24).Hydroxyl radical (HO^·) is one of the most reactive and major species generated under aerobic conditions in biological systems (1, 25, 26). Among several HO^·-mediated oxidative modifications, the protein tyrosine modification 3,4-dihydroxyphenylalanine (DOPA) has been reported as a major product and index of HO^· attack on tyrosine residues in proteins (Fig. 1) (27, 28). DOPA is also formed on protein tyrosine residues via controlled enzymatic pathways through enzymes such as tyrosinase or tyrosine hydroxylase (28). Once formed, protein-bound DOPA has the potential to initiate further oxidative reactions through binding and reducing transition metals or through redox cycling between catechol and quinone (dopaquinone) forms (29, 30). Recent studies have suggested that protein-bound DOPA is involved in triggering antioxidant defenses (30) and mediating oxidative damage to DNA (31). Moreover, elevated levels of protein-bound DOPA have been reported in several diseases, including atherosclerosis, cataracts, and myocardial disease, and in PD patients undergoing levodopa therapy (26, 32–36). However, the specific DOPA-modified proteins, which could provide mechanistic knowledge of the progression of these diseases, have not been identified (27, 28). The ability to identify site-specific protein modifications should lead to a better understanding of the role of DOPA modification in disease pathologies as well as new molecular signatures or therapeutic targets for diseases.Open in a separate windowFig. 1.DOPA and dopaquinone formation from tyrosine.Therefore, in this study, we demonstrate the ability to identify site-specific DOPA and dopaquinone (DQ) modifications on protein tyrosine residues in normal mouse brain and heart tissues and their relative stoichiometries that are present in vivo under non-stressed conditions. Such endogenous protein modifications were detected using LC-MS/MS. The results from this global proteomics survey suggests that HO^· in tissues under normal conditions is generated largely from the mitochondria and metal-binding proteins where the resulting DOPA/DQ modifications have the potential to disrupt mitochondrial respiration as well as alter tyrosine phosphorylation signaling pathways such as 14-3-3-mediated signaling in brain tissue.