Category Archives: Protein Folding

Zinc is a micronutrient and essential trace element required for metabolism reactions that catalyze number of enzyme reactions, protein folding, gene expression etc. Deficiency of zinc may cause range of infectious diseases and other complications affecting almost every aspect of health such as delayed or retarded growth in children, hair loss, lack of cognitive function, reduced sense of taste and smell, loss of appetite, immunosuppression, anemia, night blindness, etc. Zinc deficiency may also cause deficiency of vitamin A as zinc is required for vitamin A absorption. Zinc deficiency is the fifth leading factor causing disease across the globe, and according to World Health Organization (WHO) 31% of the global population is experiencing zinc deficiency.

To remain ahead of your competitors, request for a sample[emailprotected] https://www.persistencemarketresearch.com/samples/17281

Underdeveloped countries are facing major problem of death due to childhood diarrhea and pneumonia caused due to zinc deficiency. As per International Zinc Nutrition Consultative Group (IZiNCG) zinc deficiency is expected to cause 176,000 diarrhea deaths, 406,000 pneumonia deaths and 207,000 malaria deaths across the world Regions such as Africa, Middle East and South-East Asia are bearing high burden of zinc deficiency due to poor nutrition, lack of breastfeeding etc. Treatment of zinc deficiency mainly involves intake of zinc rich food and zinc supplements. Mild zinc deficiency requires treatment with zinc supplements at 2-3 times the recommended dietary allowance (RDA) and moderate to severe zinc deficiency requires zinc supplements with 4-5 times the RDA and treatment is continued for at least 6 months.

Oral repletion of zinc in the form of zinc acetate, zinc sulfate, zinc aspartate, zinc orotate and zinc gluconate, multivitamin supplements etc. and zinc supplements are available in the form of oral tablets capsules, syrups or intravenous solutions. National Institute of Health (NIH) recommends daily intake of 20-40 mg zinc for adults orally to avoid complications caused by zinc deficiency. High dose of zinc i.e. more than 50 mg per day is recommended to patients with severe zinc deficiency or patients with irreversible malabsorptive disorders. Intravenous administration of zinc is rarely recommended unless the patient is suffering from intestinal failure or on long-tern treatment with total parenteral nutrition.

To receive Methodology request here @ https://www.persistencemarketresearch.com/methodology/17281

Market for zinc deficiency treatment is primarily driven by increasing occurrence of malnutrition in underdeveloped and developing countries. Increasing incidence of anemia, hypovitaminosis A, etc. are other factors driving demand for zinc supplements across the globe. Zinc supplements are also used as an adjunctive therapy in many disorders such as alopecia, ulcers, electrolyte replenishment therapy for diarrhea, etc.; high incidence of which can propel the demand for zinc supplements over the forecast period. However, unavailability of treatment opportunities in the underdeveloped countries can be the factor which can hamper growth of global zinc deficiency treatment market.

The global market for zinc deficiency treatment is segmented on basis of product types, treatment, dosage form, distribution channel and geography:

Among treatment type, dietary supplements is expected to dominate the global market as dietary zinc supplements are recommended along with drug therapy in every patient suffering from zinc deficiency. Among all four distribution channels of zinc supplements, e-commerce is expected to experience highest growth over the forecast period.

On the basis of geography, global zinc deficiency treatment testing market is segmented into five key regions viz. North America, Latin America, Europe, Asia Pacific, and Middle East & Africa. Asia Pacific is expected to dominate the global market for zinc deficiency treatment due to high level of malnutrition in developing countries such as India, Bangladesh, Sri Lanka, etc. The region is expected to witness robust growth due to growing awareness towards malnutrition and its consequences. North America and Western Europe shows lower occurrence of zinc deficiency as adequate intake of animal food by population in the region. Despite high incidence of zinc deficiency in Middle East and Africa market growth is limited by access to zinc supplements due to low purchasing power and poor health consciousness among general population.

To receive extensive list of important regions, Request TOC here @ https://www.persistencemarketresearch.com/toc/17281

Some of the key players present in global zinc deficiency treatment market are

Link:
Demand Scenario of Zinc Deficiency Treatment Market to Remain Positive Through 2025 - Research Newspaper

INTRODUCTION

Toxin-antitoxin (TA) systems are widely distributed throughout prokaryotic genomes and have been shown to help bacteria to survive predation by bacteriophages, immune responses, and antibiotic treatments (15). In many cases, however, the roles of chromosomal TA systems remain largely unknown, primarily due to the lack of a phenotype associated with deletion mutants under in vitro laboratory conditions (69). TA systems are also widespread among mobile genetic elements, including plasmids, superintegrons, cryptic prophages, and conjugative transposons, where they contribute to their stability (10, 11).

TA systems encode two components, a toxic protein that targets an essential cellular process and an antagonistic antitoxin, which blocks toxin activity when cells are growing under favorable conditions. Although the processes that lead to toxin activation remain under debate, it has been proposed that under certain stress conditions, increased toxin transcription and synthesis may lead to activation (8, 12). This, in turn, reduces growth rate, which can provide a means to survive with minimal metabolic burden until favorable conditions return (13).

TA systems are divided into six types according to the nature of the toxin and antitoxin (whether they are RNA or protein) and the mechanism of toxin antagonism (3). Type II systems, in which a protein toxin is sequestered by a protein antitoxin, have been most extensively studied. They are also remarkably abundant in Mycobacterium tuberculosis, which potentially encodes more than 80 type II TA systems, and are thought to have contributed to the success of M. tuberculosis as a human pathogen (1416). Many of the putative M. tuberculosis toxins tested thus far were shown to inhibit bacterial growth, suggesting that these TA systems are functionally active and could modulate M. tuberculosis growth under certain conditions, thereby contributing to survival in the human host (15, 17). Accordingly, many M. tuberculosis TA operons were shown to be induced in response to relevant stressors, including hypoxia, the presence of antimicrobial drugs, or macrophage engulfment (14, 17). As M. tuberculosis encodes, among others, more than 50 VapBC, 10 MazEF, 3 HigBA, and 3 RelBE TA systems, it might be expected that there is redundancy between them, alongside condition-specific applications for each system. Furthermore, the highly toxic nature of some of these toxins suggests that their antibacterial mechanisms could be developed into antimicrobials (18).

This study focuses on a family of four putative toxins from M. tuberculosis, namely, Rv0078A, Rv0836c, Rv1045, and Rv2826c, which share a conserved nucleotidyltransferase (NTase)like domain annotated as domain of unknown function (DUF) 1814 (Fig. 1A). The most well-characterized example of this DUF1814 family is AbiEii from Streptococcus agalactiae, which shares 18.3% sequence identity with Rv1045, and was identified within the AbiE abortive infection bacteriophage-defense systems (19). AbiEii was shown to constitute a new type of TA system, type IV, based on the observation that no interaction could be detected between the toxin and the antitoxin proteins (20). The DUF1814 family of proteins is widespread in bacterial, archaeal, and fungal genomes (20), though not all examples are genetically linked to putative antitoxins. As putative NTases, DUF1814 proteins contain four conserved motifs. The N-terminal motifs I and II are found in DNA polymerase and are proposed to coordinate a metal ion for nucleotide binding and transfer (20). The C-terminal motif III is similar to that of tRNA NTases that add the 3-CCA motif to immature tRNAs and may be important for base stacking with substrates (21). The C-terminal motif IV is unique to DUF1814 proteins and is proposed to form a catalytic site with motif III (20).

(A) Scaled representation of the four M. tuberculosis TA systems containing NTase-like toxin genes with original and revised nomenclature (left), and corresponding toxicity and antitoxicity assays in M. smegmatis (right). For toxicity and antitoxicity assays, cotransformants of M. smegmatis mc2 155 containing pGMC-vector, -MenT1, -MenT2, -MenT3, or -MenT4 (toxins) and pLAM-vector, -MenA1, -MenA2, -MenA3, or -MenA4 (antitoxins) were plated on LB-agar in the presence or absence of anhydrotetracycline (Atc; 100 ng ml1) and acetamide (Ace; 0.2%) inducers for toxin and antitoxin expression, respectively. Plates were incubated for 3 days at 37C. T and A denote toxin and antitoxin, respectively. and + represent absence or presence of inducer, respectively. (B) M. smegmatis strain mc2 155 transformed with plasmid pGMCS-TetR-P1-RBS1-MenT3 was grown in complete 7H9 medium with Sm. At time 0, the culture was divided into two. Half was kept in the same medium (pale blue bars) and half was additionally treated with Atc (200 ng ml1) (dark blue bars). Samples were harvested at the indicated times, washed, diluted, and plated on LB-agar with Sm but without Atc. Colonies were counted after 3 days at 37C. Shown values are the average of three biological replicates with SD. CFU, colony-forming unit. (C) Samples of the same cultures as in (B) were harvested after 8 or 24 hours, labeled with the LIVE/DEAD BacLite dyes [Syto 9; propidium iodide (PI)], and analyzed by fluorescence-activated cell sorting. The percentage of PI-positive cells is shown for each sample (pale blue bars, no Atc; dark blue bars, 200 ng ml1 Atc). Shown values are the average of three biological replicates with SD. (D) M. tuberculosis wild-type (WT) H37Rv or mutant strain H37Rv (menA3-menT3)::dif6 were transformed with 100 ng of plasmids expressing either menA3, menT3, or menA3-menT3. These plasmids encode a consensus Shine-Dalgarno sequence (RBS1), except for Weak-RBS-menT3, which encodes a near-consensus sequence (RBS4) to weaken expression. After phenotypic expression, half of the transformation mix was plated on 7H11 oleic acidalbumin-dextrose-catalase (OADC) plates with Sm, and the other half was plated on 7H11 OADC Sm plates supplemented with Atc (200 ng ml1). Plates were imaged after 20 days at 37C; data are representative of three independent experiments. (E) Mutant strain H37Rv (menA3-menT3)::dif6 was transformed with 100 ng of plasmids expressing either menT3 WT or mutant alleles introducing the D80A, K189A, or D211A substitutions. After phenotypic expression, half of the transformation mix was plated on 7H11 OADC plates with Sm, and the other half was plated on 7H11 OADC Sm plates supplemented with Atc (200 ng ml1). Pictures were taken after 20 days at 37C; data are representative of three independent experiments.

In M. tuberculosis, the DUF1814 toxins are encoded downstream of a variety of putative antitoxins (Fig. 1A). The toxin gene rv0078A is paired with a short upstream open reading frame encoding a 68amino acid antitoxin, Rv0078B, related to MazE antitoxins, which is predicted to be disordered and lacking a DNA-binding domain (16). Toxin gene rv0836c lies downstream of a COG4861 gene, encoding a much larger putative antitoxin than the cognate toxin (Fig. 1A). Rv1045 and Rv2826c toxins are downstream of their cognate putative antitoxins Rv1044 and Rv2827c, respectively, both of which are COG5340 transcriptional regulator family proteins (Fig. 1A). COG5340 proteins include the S. agalactiae AbiEi antitoxin partner of AbiEii, which has previously been shown to bind to and repress the abiE promoter, similar to autoregulation observed in type II TA loci (22). An earlier transposon site hybridization study identified both the Rv1044 and Rv2827c antitoxins as essential for growth (23). Saturating transposon mutagenesis has additionally demonstrated that Rv1044 is essential, while transposon insertions in Rv2827c impart a growth defect (24). The fact that both antitoxins are important for M. tuberculosis growth strongly suggests that their putative cognate Rv1045 and Rv2826c toxins inhibit growth in M. tuberculosis.

Here, we undertook a series of microbiological, structural, genetic, and biochemical studies to investigate the DUF1814 toxins of M. tuberculosis and reveal their mode of action. We show that the Rv1045 toxin is a tRNA NTase that is active in M. tuberculosis and blocks translation through a previously undescribed mechanism involving inactivation of serine tRNAs.

We first investigated the activity of the putative TA systems containing NTase-like DUF1814 toxins in Mycobacterium smegmatis, which is closely related to M. tuberculosis and does not encode similar antitoxins (15). On the basis of the findings presented below, we renamed these putative systems as mycobacterial AbiE-like NTase toxins (MenT) and antitoxins (MenA), numbered according to their order in the M. tuberculosis genome (Fig. 1A). Toxins and antitoxins were expressed in trans, with the toxins cloned into the pGMC-integrative plasmid under the control of an anhydrotetracycline (Atc)inducible promoter and the antitoxins into the compatible pLAM plasmid under the control of an acetamide (Ace)inducible promoter (Fig. 1A). Among the four putative toxins, only MenT1 has been tested so far and was shown to be toxic in M. smegmatis when expressed without the upstream open reading frame encoding MenA1, suggesting that MenA1-MenT1 form a functional TA system (16). Accordingly, the data presented in Fig. 1A show that MenT1 toxicity was efficiently counteracted by MenA1 expressed in trans. Both MenA3-MenT3 and MenA4-MenT4 also acted as TA pairs, while MenT2 expression was not toxic (Fig. 1A). Inhibition of MenT4 toxicity could only be achieved when the putative antitoxin was expressed in the context of the menA4-menT4 operon (Fig. 1A). Expression of MenA4 alone from pLAM was toxic (fig. S1A), indicating that MenA4-MenT4 might not function as a typical TA pair under these conditions. Similar experiments performed in Escherichia coli confirmed the phenotypes observed in M. smegmatis for MenA2-MenT2, MenA3-MenT3, and MenA4-MenT4 (including MenA4 toxicity), but not for MenT1, which exhibited no detectable toxicity in E. coli (fig. S1B). Last, coexpression of the active toxins with noncognate antitoxins did not reveal any detectable cross-talk between the different TA pairs (fig. S1, A and C). Note that cross-talk assays with MenA4 antitoxin expressed from pLAM in M. smegmatis could not be performed because of its toxicity.

Ectopic expression of MenT3 in the presence of inducer showed the most robust toxicity in both M. smegmatis and E. coli when compared to the other toxins (Fig. 1A and fig. S1). In M. smegmatis, only a few MenT3 transformants were obtained, even in the absence of inducer. Ectopic expression of MenT3 in M. smegmatis induced a rapid drop of about 3-log10 in colony-forming units only 2 hours after induction with Atc (Fig. 1B). LIVE/DEAD BacLight stains have previously been used to study the effects of toxin expression on cell viability in M. tuberculosis (18). Flow cytometry analysis of M. smegmatis expressing MenT3 revealed that the proportion of propidium iodidepermeable cells was substantially higher in MenT3-induced versus noninduced cells 8 or 24 hours after induction with Atc (Fig. 1C), indicating that MenT3 strongly affects cell viability.

To investigate the impact of MenA3 and MenT3 on M. tuberculosis growth, plasmids encoding the toxin, the antitoxin, or both, were introduced into H37Rv wild-type (WT) strain. The resulting transformants were not sensitive to ectopic expression of MenT3 (Fig. 1D), presumably because endogenous MenA3 was sufficient to neutralize the sum of endogenous and ectopic MenT3. To confirm this hypothesis, we attempted to construct a strain deleted for the menA3-menT3 operon. Previous work showed that menA3 cannot be disrupted by transposon insertion (24). Accordingly, we found that deletion of the menA3-menT3 operon in M. tuberculosis H37Rv strain could not be achieved, most likely because simultaneous disruption of both genes resulted in a toxic effect from residual MenT3. To circumvent this problem, we constructed the deletion in a derivative of H37Rv carrying a second copy of menA3 constitutively expressed from a pGMC integrative plasmid. Once the menA3-menT3 operon was deleted, it was then possible to remove the ectopic copy of menA3 by pGMC plasmid replacement (fig. S2). The menA3-menT3 mutant became highly sensitive to the MenT3 toxin, even in the absence of inducer (Fig. 1D). Therefore, to finally obtain transformants, menT3 was cloned downstream of a weaker Shine-Dalgarno sequence. Using this construct, we observed inducible MenT3 toxicity, which was fully abolished by the presence of the antitoxin (Fig. 1D). Together, these data demonstrate that the MenT3 toxin inhibits growth and that MenA3-MenT3 functions as a bona fide TA pair in M. tuberculosis.

A previous amino acid sequence alignment of DUF1814 putative NTases highlighted conserved residues, a number of which were confirmed as essential for AbiEii toxicity in S. agalactiae (20). To investigate whether some of these residues were important for MenT3 toxicity, we selected and engineered three conserved residues for substitution: D80A, localized in the DNA pol superfamily motif, and K189A and D211A, both toxin-specific residues. We then tested the impact of these substitutions on M. tuberculosis growth (Fig. 1E). All three substitutions abolished MenT3 toxicity in both M. tuberculosis (Fig. 1E) and E. coli (fig. S3A).

Next, we investigated whether the MenT3 toxin and MenA3 antitoxin could interact in vivo. Since this TA pair is functional in E. coli, we performed affinity-tagged in vivo copurification experiments in E. coli using His-tagged variants of MenT3 and MenA3, which were first confirmed to be active as toxin and antitoxin, respectively (fig. S4A). In strains coexpressing both the toxin and the antitoxin (with either the toxin or the antitoxin tagged), and with tagged toxin and tagged antitoxin alone as controls, the in vivo copurification revealed that a small but significant fraction of the MenT3 toxin and the MenA3 antitoxin copurified, whether the toxin or the antitoxin was used as bait (fig. S4, C and D). Similar results were obtained with the MenA1-MenT1 pair, which encodes a much shorter, unrelated antitoxin (fig. S4, B, E, and F). Together, these data show that both TA partners can interact, but it remains to be determined whether a direct interaction between an NTase toxin and its cognate antitoxin is required for toxin inhibition.

To begin investigations into the mechanism of toxicity of MenT3, we solved its structure to 1.6 resolution by x-ray crystallography (Fig. 2A and Table 1). MenT3 is a monomeric bi-lobed globular protein, with two hemispheres connected by a short linker (Fig. 2A). This monomeric assembly matches the expected size observed by size exclusion chromatography. Surface electrostatics show a distinct electropositive surface leading to a deeper recess (Fig. 2B, left), which contains residues D80, K189, and D211 that were needed for toxicity in vivo (Fig. 1E). This potentially indicates the position of the active site, and the electropositive surface may facilitate interaction with electronegative substrates such as nucleic acids. To further characterize the DUF1814 family, we also solved the MenT4 toxin structure to 1.2 resolution (Fig. 2C and Table 1). MenT4 is also monomeric (also observed by size exclusion chromatography) and the overall architecture is similar to, but not exactly the same as, MenT3. MenT4 has a bi-lobed globular structure and distinct electropositive patches close to a similarly positioned active site region (Fig. 2, C and D). Aligning MenT3 and MenT4 by sequence gave a poor root mean square deviation (RMSD) of 13.4 ; however, this can be improved to 4.7 using sequence-independent superposition, which demonstrates similarity in overall fold (Fig. 2E). A close-up of MenT3 residues D80, K189, and D211 show them clustered at the putative active site, and when overlaid, the homologous MenT4 residues, D69, K171, and D186, respectively, take up similar positions (Fig. 2F). There was also density for a phosphoserine at MenT3 S78, but the corresponding residue in MenT4, S67, was not phosphorylated (Fig. 2F). Searches for structural homologs of MenT3 and MenT4 were performed using the DALI server (25). Among multiple hits for NTases, the best match was for JHP933 from Helicobacter pylori, a predicted NTase encoded by the jhp0933 gene (26). JHP933 aligned to MenT3 with an RMSD of 2.4 , though multiple additional helices were resolved in the MenT3 structure (Fig. 2G). An analysis of the H. pylori genome revealed that the jhp0932 gene lies just upstream of jhp0933 and partially overlaps its coding sequence. The presence of these genes in what appears to be a classic TA configuration suggests that JHP933 may belong to the MenT3/MenT4 family of NTase-like toxins.

(A) Structure of monomeric MenT3 toxin, with views from front and back, shown as cyan cartoon representations. (B) Surface electrostatics of MenT3, viewed as in (A), with red for electronegative and blue for electropositive potential. (C) Structure of monomeric MenT4, with views from front and back, shown as salmon cartoon representations. (D) Surface electrostatics of MenT4, viewed as in (C), colored as per (B). (E) Superposition of MenT4 onto MenT3, viewed and colored as per (A) and (C). (F) Tilted close-up view of the toxin active sites, as indicated by the boxed region of (E). MenT3 residues S78 (phosphorylated), D80, K189, and D211 are indicated, along with the homologous MenT4 residues S67, D69, K171, and D186. (G) Alignment of JHP933 (PDB: 4O8S) as orange cartoon representation, against MenT3 viewed and colored as per (A, left).

MenT3 is the most toxic of the four M. tuberculosis NTase-like toxins tested, both in mycobacteria and in E. coli (Fig. 1 and fig. S1). We therefore took advantage of this robust toxicity to search for E. coli genes that were able to suppress MenT3-mediated growth inhibition when overexpressed. We reasoned that identification of such suppressors might potentially shed light on the cellular processes affected by the toxin. Details of the genetic selection used are described in Materials and Methods. Among the approximately 60,000 clones of the E. coli genomic plasmid library tested in this work, we identified 18 plasmids that passed two rounds of selection and appeared to encode bona fide suppressors of MenT3 toxicity. We observed that the toxin-resistant colonies were noticeably smaller and translucent compared to noninduced cells, indicating that, although notably reduced, MenT3 toxicity is not fully suppressed. Sequencing of the genomic regions encoded by the 18 suppressor plasmids revealed that several of these candidate plasmids harbored the same genomic fragments. Six different suppressor clones encompassing two different regions of the E. coli chromosome were identified. Two of the six suppressor plasmids harbored the ydeA gene, encoding an l-arabinose (l-ara) exporter protein known to decrease l-ara levels in E. coli (27). These suppressors were discarded as YdeA overexpression would presumably decrease toxicity of many toxic proteins expressed from the araBAD promoter. The four other suppressor plasmids harbored the rph gene, encoding the phosphorolytic ribonuclease (RNase PH), involved in the 3 processing of RNAs (Fig. 3A). RNase PH removes nucleotides downstream of the 3-CCA sequence, required for aminoacylation of tRNAs, from tRNA precursors with 3 extensions. It is also involved in other RNA maturation and quality control processes, including the maturation of rRNA (28).

(A) The E. coli K-12 genomic region containing the rph gene is shown. Suppressor plasmids that counteract MenT3 toxicity encoded rph, as depicted by small arrows under the adjacent genes pyrE, yicC, and dinD. The positions in base pair of the ends of each suppressor fragment, in relation to the E. coli K-12 chromosome, are indicated between brackets. (B) Overexpression of E. coli RNase PH partially suppresses MenT3 toxicity. E. coli DLT1900 strains containing either pK6-vector () or pK6-MenT3 (+) were cotransformed with p29SEN-vector () or p29SEN-Rph (RNase PH) (+). The resulting cotransformants were serially diluted, spotted onto LB-agar plates in the presence or absence of l-ara (0.1%) and IPTG (200 M) inducers, and incubated at 37C. (C) Deletion of rph further increases MenT3 toxicity. Transformants of E. coli DLT1900 WT and rph mutant strains containing plasmid pK6-MenT3 were serially diluted, spotted onto LB-agar plates with or without l-ara (0.01%), and incubated at 37C. (D) In vitro transcription/translation reactions assessing levels of DHFR control protein produced in the absence or presence of increasing concentrations of MenT3 toxin. Samples were separated by SDSpolyacrylamide gel electrophoresis and stained with InstantBlue. (E) For in vivo assays, transformants of E. coli BL21 (DE3) containing plasmid pET-MenT3 or the empty vector were grown in M9M at 37C. Following overexpression of MenT3, tRNAs were extracted, separated, and visualized by Northern blot using specific radiolabeled probes against tRNATrp. For in vitro assays, purified MenT3 (10 M) was added to transcription/translation assays producing GatZ protein. After 2 hours at 37C, tRNAs were extracted, separated, and visualized by Northern blot as performed for the in vivo samples. All images are representative of triplicate data.

Suppression of MenT3 toxicity by RNase PH overexpression was confirmed by cloning rph alone in a lowcopy number plasmid under the control of an isopropyl--d-thiogalactopyranoside (IPTG)inducible promoter and assaying for growth in the presence of MenT3 in E. coli (Fig. 3B). We also showed that the toxicity of MenT3 was enhanced when expressed in E. coli carrying a deletion of the rph gene, even with a 10-fold decrease in inducer levels (Fig. 3C), further reinforcing the genetic link between menT3 and rph. The primary role of RNase PH in processing tRNAs suggests that DUF1814 NTase-like toxins could act directly at the site of aminoacylation at the 3-end of tRNA, thus inhibiting translation. Whether endogenous RNase PH would be sufficiently induced in response to toxin expression to help restore the functional tRNA pool in recovering M. tuberculosis cells remains to be determined.

MenT3 WT and the MenT3(D80A) and MenT3(K189A) substitutions were overexpressed and purified for biochemical characterization. When tested in an in vitro transcription/translation reaction that uses recombinant E. coli components, purified MenT3 WT reduced production of the E. coli dihydrofolate reductase (DHFR) control protein in a concentration-dependent manner (Fig. 3D). Compared to MenT3 WT, MenT3(D80A) and MenT3(K189A) had a markedly reduced impact on the production of DHFR (fig. S5A). The same trend was observed when MenT3 WT, MenT3(D80A), and MenT3(K189A) were used in in vitro reactions producing WaaF and GatZ as test proteins (fig. S5, B and C). We also expressed and purified MenT4 WT and demonstrated that this, too, prevented the production of DHFR in a concentration-dependent manner in in vitro transcription/translation assays (fig. S5D).

The fact that MenT3 inhibited protein synthesis, and that RNase PH is involved in the removal of nucleotides following the 3-CCA sequence required for tRNA aminoacylation, suggested that tRNA charging might be affected by MenT3 expression in vivo. To address this hypothesis, we first used a method developed for E. coli, which separates charged from uncharged tRNAs and allows their detection by Northern blot after extraction in vivo (29). We chose tRNATrp as a model tRNA because (i) the tryptophanyl-tRNA can be well separated from uncharged tRNATrp and (ii) there is only one tRNATrp in E. coli (29). No charged tryptophanyl-tRNATrp could be detected following overexpression of MenT3 when compared to the empty vector control (Fig. 3E and fig. S5E). tRNATrp charging levels were also investigated in vitro by adding purified MenT3 to the transcription/translation assay described above (Fig. 3E). In this case, MenT3 also affected tRNATrp charging in vitro, thus supporting the hypothesis that the toxin inhibits protein synthesis by preventing aminoacylation of tRNA.

The observation that MenT3 is related to NTases (Fig. 2G) suggests that its mode of action is to directly transfer nucleotides to tRNAs, thereby preventing aminoacylation. We performed assays using radiolabeled tRNAs to track the addition of nucleotides by MenT3 WT, MenT3(D80A), and MenT3(K189A) (Fig. 4).

(A) Radiolabeled E. coli tRNATrp was incubated with 1, 0.1, 0.01, or 0.001 g of MenT3 WT or no toxin () for 20 min at 37C in the presence of unlabeled GTP, ATP, UTP, or CTP. Extended products are indicated with arrowheads throughout all panels. (B) Radiolabeled E. coli tRNATrp was incubated with 1, 0.1, or 0.01 g of MenT3 WT or MenT3(D80A) with CTP or UTP, as per conditions in (A). (C) Incubation of radiolabeled E. coli tRNATrp with 1, 0.1, 0.01, or 0.001 g of MenT3 WT or MenT3(K189A), with CTP, UTP, or a mixture of both, as per conditions in (A). (D) Radiolabeled E. coli tRNATrp preparations, made with or without a 3-CCA motif, were incubated with 1, 0.1, or 0.01 g of either MenT3 WT, MenT3(K189A), or no toxin (), for 20 min at 37C in the presence of unlabeled UTP or CTP. Note that the () CCA lanes have been overexposed to equalize intensity to the (+) CCA lanes of the same gel. Assays of the individual WT and MenT3 substitution proteins and tRNATrp CCA substrates shown in (A) to (D) were performed between two and four times.

MenT3 WT was incubated with tRNATrp from E. coli, as a model recipient tRNA, in the presence of guanosine 5-triphosphate (GTP), adenosine 5-triphosphate (ATP), uridine 5-triphosphate (UTP), or cytidine 5-triphosphate (CTP), and nucleotide transfer was monitored as an increase in tRNA size by high-resolution polyacrylamide gel electrophoresis (PAGE; Fig. 4A). At high concentrations of the enzyme, we found that MenT3 can add two to three extra nucleotides to tRNATrp in the presence of CTP or UTP, with a slight preference for CTP, suggesting that MenT3 is a pyrimidine-specific NTase (Fig. 4A). No transfer was observed with purines ATP or GTP as substrates (Fig. 4A). MenT3(D80A), which was unable to inhibit in vitro protein synthesis (fig. S5, A to C), had no NTase activity with either UTP or CTP (Fig. 4B). MenT3(K189A), which was also inactive in the in vitro transcription/translation assay (fig. S5, A to C), only lost its NTase activity in the presence of UTP, but retained some activity (albeit less than WT) in the presence of CTP, or both nucleotides (Fig. 4C). This could imply that K189A is important for substrate nucleotide selectivity. No synergistic effect was seen when MenT3 WT was incubated with a mixture of CTP and UTP, as the pattern with both nucleotides together resembled that of CTP alone (Fig. 4C).

Canonical tRNA NTases typically add the 3-CCA motif to tRNAs lacking an encoded 3-CCA that are processed at the level of the discriminator nucleotide (nucleotide 73). They also repair this motif when 3-exoribonucleases, such as RNase PH, fail to stop at the 3-CCA motif when processing tRNA precursors containing an encoded 3-CCA, typically removing the terminal A residue. Since M. tuberculosis contains a mixture of tRNA genes encoding or lacking a 3-CCA motif, we wondered whether MenT3 had a preference for one class (or another class) of substrate. While faint NTase activity was observed when MenT3 WT and MenT3(K189A) were incubated with CTP and tRNATrp lacking a 3-CCA, the data show that MenT3 had a clear preference for tRNAs that already possessed a 3-CCA motif (Fig. 4D). This is in contrast to the normal function of tRNA NTases, which prefer tRNAs lacking an intact 3-CCA. Again, MenT3 WT modified tRNATrp using both CTP and UTP as substrate, while MenT3(K189A) could only use CTP (Fig. 4D). Addition of nucleotides to mature tRNAs by MenT3 would completely abolish the ability of these tRNAs to be charged with their cognate amino acid and take part in translation, accounting for their cellular toxicity.

Our in vivo data show that toxins MenT3, as well as MenT1 and MenT4, are significantly less toxic in E. coli than in mycobacteria (Fig. 1 and fig. S1), which suggests that these toxins may have a tRNA target preference. We therefore asked whether MenT3 would exhibit some specificity toward the different tRNAs of M. tuberculosis. We made polymerase chain reaction (PCR) templates allowing us to in vitro transcribe the 45 different tRNAs of M. tuberculosis, each with a 3-CCA motif (fig. S6). As before, each radiolabeled tRNA was incubated with MenT3 and nonradiolabeled CTP (Fig. 5A). To our surprise, MenT3 appeared to be highly specific, preferentially modifying the four M. tuberculosis tRNASer isoacceptors, along with weak modification of tRNALeu5 (Fig. 5A). Although we cannot exclude that MenT3 can modify other tRNAs in vivo, the data show that the toxin presents a high degree of specificity toward different tRNAs in vitro, which may explain the variable toxicity observed in different bacteria.

(A) Radiolabeled M. tuberculosis tRNAs were incubated with 0.1 g of MenT3 WT (+) or no toxin () for 20 min at 37C in the presence of unlabeled CTP. E. coli tRNATrp (EcTrp) was used as a positive control. The global screen of all M. tuberculosis tRNA was performed once and the effect of MenT3 tRNASer2 was confirmed twice independently. (B) Schematic diagram of the MenT3 toxin mechanism of action. MenT3 elongates the 3-CCA motif of specific tRNAs, preventing their charging by aminoacyl-tRNA synthetases (AaRS), thereby interfering with translation and inhibiting bacterial growth.

Last, we asked whether the antitoxin MenA3 inhibited the NTase activity of MenT3 directly, or whether it could simply reverse its action by removing the added nucleotides in a manner similar to the RNase PH multicopy suppressor. Addition of MenA3 strongly inhibited the NTase activity of MenT3 on the natural substrate M. tuberculosis tRNASer2 when coincubated with the toxin at a molar ratio > 2.5. However, MenA3 failed to remove the added nucleotides from tRNASer2 when added after a preincubation of tRNASer2 with MenT3, even at high concentrations (fig. S7). This suggests that the antitoxin is likely to inhibit the toxin rather than reverse the reaction on the substrate.

This study has characterized a family of TA systems from M. tuberculosis containing NTase-like DUF1814 toxins, establishing MenT3 as a potent toxin in this problematic pathogen. We have solved the structures of the homologous toxins MenT3 and MenT4 by x-ray crystallography, revealing fold similarity and conserved residues within the proposed active sites, and have observed a similar mode of toxin activity, targeting protein synthesis. We have further elucidated the mechanism of toxicity for MenT3, showing that it functions as a pyrimidine-specific NTase preferentially targeting M. tuberculosis tRNASer in vitro (Fig. 5B).

The observation that the three NTase toxins identified in this work show different levels of toxicity when expressed in the same host, and that such toxic signatures can vary when expressed in different bacterial hosts (i.e., E. coli versus mycobacteria), is intriguing (Fig. 1 and fig. S1). The most marked example is MenT1, which shows robust toxicity in M. smegmatis but no toxicity in E. coli (Fig. 1A and fig. S1B). Although we cannot exclude this being a result of improper folding or expression of the toxin in E. coli, it is also reasonable to assume that the toxin may not be able to recognize its tRNA targets due, for example, to tRNA modification, or the absence of its preferred tRNA target (30). Another possibility is that tRNA targets are expressed at higher levels in E. coli and are thus sufficiently abundant to overcome the noxious effect of the toxin in vivo. The fact that M. tuberculosis and M. smegmatis only have 45 and 46 tRNA genes, respectively, while E. coli has 86, is in line with this hypothesis (30, 31).

The apparent in vitro specificity of MenT3 for certain M. tuberculosis tRNAs, especially tRNASer, is remarkable (Fig. 5A). We did not test other tRNAs in E. coli besides tRNATrp; it may well have been fortuitous that the only tRNA we tested in this organism was detectably modified by the toxin in vitro (Fig. 4A) and in vivo, inferred from the reduced charging levels following toxin expression (Fig. 3E). We checked whether the M. tuberculosis tRNAs that were substrates of MenT3 had any distinguishing features and were struck by the fact that all serine tRNAs and several leucine tRNAs were unique among M. tuberculosis tRNAs in that they had long variable arms (fig. S8A) (32). While this is intriguing and may contribute to substrate specificity, it cannot be the only recognition element because (i) two leucine tRNAs besides tRNALeu5 have variable loops but are not MenT3 substrates in vitro and (ii) E. coli tRNATrp does not have a variable loop (fig. S8B), but can be extended by the NTase activity of the toxin. It is also intriguing in this regard that M. tuberculosis tRNATrp is not a MenT3 substrate in vitro. E. coli and M. tuberculosis tRNATrp are highly homologous but do show differences in their variable- and T-arm sequences (fig. S8C). Substrate specificity therefore appears to come from a combination of multiple sequence and structure motifs. Having identified these tRNA targets in vitro, further work is now needed to confirm targeting in vivo in M. tuberculosis.

Our observed TA interactions raise questions regarding the molecular mechanisms of antitoxicity for DUF1814 toxins (fig. S4, C to F). Typically, in type II TA systems, antitoxin function is in part driven by its strong and direct interaction with the cognate toxin (3). While we have shown interactions between cognate toxins and antitoxins (fig. S4, C to F), the antitoxin interaction in vivo appears weak. We additionally demonstrated that coincubation of the MenA3 antitoxin with MenT3 is able to neutralize the NTase activity (fig. S7). This suggests that any interaction-based antitoxicity might be a transient and labile mechanism and, due to the difference in size and sequence between antitoxins MenA1 and MenA3 (Fig. 1A), may well differ between these systems.

The DUF4433 DarT toxin from M. tuberculosis was recently identified as a single-stranded DNA NTase that specifically and reversibly adenosine 5-diphosphate (ADP)ribosylates thymidines (33). Our study identifies MenT3 as an NTase toxin from the unrelated DUF1814 protein family. In comparison to DarT, MenT3 acts via a distinct and novel mode of toxicity where the MenT3 toxin preferentially targets M. tuberculosis tRNAs in vitro, preventing their charging with cognate amino acids by adding nucleotides to the 3-CCA acceptor stem (Fig. 5B). Accordingly, antitoxin function also appears to differ between these systems. Whereas DarT is counteracted enzymatically by the cognate antitoxin DarG via target de-ADP-ribosylation (33), we found that MenA3 was unable to reverse MenT3 toxicity by removing nucleotides, suggesting that MenA3 likely inhibits the toxin activity.

Increasing numbers of toxins have been identified that target tRNAs by various mechanisms (13). The M. tuberculosis type II VapC toxins function as endoribonucleases cleaving tRNAs (34), whereas TacT from Salmonella Typhimurium and AtaT from E. coli are tRNA acetyltransferases, modifying charged tRNAs to block translation (35, 36). That MenT3 provides yet another way to inhibit tRNA activity is perhaps not unusual, given the essential nature of translation to cellular growth and survival. This likely reflects the value of possessing multiple TA systems to promote adaptability to different stressful environments via tRNA metabolism, with downstream effects ranging from stalling cell growth to potentially altering translation output (13). It remains to be seen whether this mechanism is conserved among DUF1814-toxins; while MenT4 shares structural similarities to MenT3 and inhibits protein synthesis in vitro (Fig. 2 and fig. S5D), we have not yet explored the molecular mechanism behind its toxicity. Given the continued significance of M. tuberculosis worldwide, the mechanism used by the MenA3-MenT3 TA system highlights a new way to block protein synthesis. We propose that further exploring the molecular mechanisms of both toxicity and antitoxicity will provide useful insights into the regulation of bacterial growth.

E. coli DH5 (Invitrogen), DH10B (Thermo Fisher Scientific), BL21 (DE3) (Novagen), ER2566 (New England Biolabs), W3110 [strain American Type Culture Collection (ATCC) 27325], DLT1900 (37), and M. smegmatis mc2 155 (strain ATCC 700084) are as previously described. To construct BL21 (DE3) slyD, the slyD::KmR allele from JW3311 (Keio collection) was moved into BL21 (DE3) using bacteriophage P1-mediated transduction. To construct the unmarked DLT1900 rph mutant, the rph::KmR allele from JW3618 (Keio collection) was first moved into DLT1900 using bacteriophage P1-mediated transduction and by subsequent removing of the kanamycin (Km) resistance cassette using plasmid pCP20, as previously described (38). E. coli were routinely grown at 37C in LB medium or M9 minimal (M9M) medium supplemented when necessary with Km (50 g ml1), ampicillin (Ap; 50 g ml1), chloramphenicol (Cm; 34 g ml1), streptomycin (Sm; 25 g ml1), spectinomycin (Sp; 50 g ml1), IPTG (1 mM), l-ara (0.1% w/v), or d-glucose (glu; 0.2% w/v). M. smegmatis mc2 155 strains were routinely grown at 37C in either LB or 7H9 medium (Difco). M. tuberculosis H37Rv (WT; ATCC 27294) and mutant strains were routinely grown at 37C in complete 7H9 medium (Middlebrook 7H9 medium, Difco) supplemented with 10% albumin-dextrose-catalase (ADC; Difco) and 0.05% Tween 80 (Sigma-Aldrich), or on complete 7H11 solid medium (Middlebrook 7H11 agar medium, Difco) supplemented with 10% oleic acidADC (OADC; Difco). When required, mycobacterial growth media were supplemented with Km (50 g ml1), hygromycin (Hm; 50 g ml1), Sm (25 g ml1), zeocin (Zc; 25 g ml1), Ace (0.2% w/v), or Atc (100 or 200 ng ml1).

Plasmids pMPMK6 (39), p29SEN (40), pGMCS (41), pGMCZ (42), pLAM12 (43), pETDuet-1, pET15b and pRARE (Novagen), pBAD30 (44), and pTA100 (4) have been described. Primers used for plasmid construction are described in table S1. All the plasmids constructed in this work have been verified by sequencing. The pMPMK6 derivatives expressing the toxins, namely, pK6-MenT1, pK6-MenT2, pK6-MenT3, and pK6-MenT4, were constructed as follows: menT1, menT2, menT3, and menT4 were PCR-amplified from the M. tuberculosis H37Rv genome and cloned as Eco RI/Hind III fragments (menT1 and menT2) and Mfe I/Hind III fragments (menT3 and menT4) into Eco RI/Hind IIIdigested pMPMK6.

The p29SEN plasmid derivatives encoding the antitoxins, namely, p29SEN-MenA1, p29SEN-MenA2, p29SEN-MenA3, and p29SEN-MenA4, were constructed as follows: menA1, menA2, menA3, and menA4 were PCR-amplified from the M. tuberculosis H37Rv genome and cloned either as Eco RI/Hind III fragments (menA1, menA2, and menA3) or as Mfe I/Hind III fragments (menA4) into Eco RI/Hind IIIdigested p29SEN. For p29SEN-Rph, the rph gene was PCR-amplified from the E. coli DLT1900 genome and cloned as an Eco RI/Hind III fragment into Eco RI/Hind IIIdigested p29SEN.

To construct pGMC-MenT2, pGMC-MenT3, and pGMC-MenT4, menT2, menT3, and menT4 were PCR-amplified using pK6-MenT2, pK6-MenT3, and pK6-MenT4 templates, respectively, and cloned into pGMCS using In-Fusion HD Cloning Kits (Takara Bio). Plasmid pGMC-MenT1 and pGMC-MenT1-His were obtained following PCR amplification of menT1 and menT1-His using pK6-MenT1 as a template and homologous recombination in linearized pGMCS plasmid by In-Fusion HD Cloning Kits (Takara Bio). For pGMC-*MenA4-MenT4, the menA4-menT4 operon was PCR-amplified from the H37Rv genome and cloned into linearized pGMCS plasmid by In-Fusion HD Cloning Kits (Takara Bio).

To construct plasmids pLAM-MenA2, pLAM-MenA3, and pLAM-MenA4, menA2, menA3, and menA4 were PCR-amplified using p29SEN-MenA2, p29SEN-MenA3, and p29SEN-MenA4 as templates, respectively. These were cloned as Nde I/Eco RI fragments (menA2 and menA3) and Nde I/Mfe I fragments (menA4) into Nde I/Eco RIdigested pLAM12. Plasmid p29SEN-MenA1 was used to amplify menA1 and menA1-His, which were then cloned as Nde I/Eco RI fragments into Nde I/Eco RIdigested pLAM12 to produce pLAM-MenA1 and pLAM-MenA1-His, respectively.

The pET vector derivatives used in this work were constructed as follows. To construct plasmid pET-MenT3-His, menT3-His (with an added fragment encoding a Ser-Ser-Gly-His6 C-terminal tag) was PCR-amplified from pK6-MenT3 template and cloned as an Nde I/Mfe I fragment into Nde I/Mfe Idigested pETDuet-1. Plasmid pET-MenT3-His was used as a template to construct pET-MenT3-His(D80A) and pET-MenT3-His(K189A) by QuikChange site-directed mutagenesis (Agilent) using appropriate primers. Plasmid pET-MenA3-His, encoding an N-terminal His6-tagged MenA3 antitoxin, was constructed by PCR amplification of menA3-His using p29SEN-MenA3 as a template, Nde I/Hind III digestion, and cloning into Nde I/Hind IIIdigested pET15b plasmid. To construct plasmid pET-MenT3/MenA3-His, menA3-His was first PCR-amplified from p29SEN-MenA3 template and cloned as an Nco I/Hind III fragment into Nco I/Hind IIIdigested pETDuet-1. menT3 was then PCR-amplified from pK6-MenT3, digested with Nde I/Mfe I, and cloned into Nde I/Mfe Idigested pET-MenA3-His. To construct pET-MenT3-His/MenA3, menA3 was first PCR-amplified using p29SEN-MenA3 as a template and cloned as an Nco I/Hind III fragment into Nco I/Hind IIIdigested pET-MenT3-His. To generate pET-MenT1-His (expressing MenT1 with an N-terminal His6-Ser-Ser-Gly-tag), menT1-His was PCR-amplified from pK6-MenT1 and cloned as an Nde I/Mfe I fragment into Nde I/Mfe Idigested pETDuet-1. For pET-MenA1-His (expressing MenA1 with an N-terminal His6-Ser-Ser-Gly-tag), menA1-His was PCR-amplified from p29SEN-MenA1 template and cloned as an Nco I/Bam HI fragment into Nco I/Bam HIdigested pETDuet-1. For pET-MenT1/MenA1-His, menT1 was PCR-amplified from pK6-MenA1 and cloned as an Nde I/Mfe I fragment into Nde I/Mfe Idigested pET-MenA1-His. For pET-MenT1-His/MenA1, menA1 was PCR-amplified from p29SEN-MenA1 and cloned as an Nco I/Bam HI fragment into Nco I/Bam HIdigested pET-MenT1His.

To generate MenT3 and MenT4 expression constructs for crystallization and biochemistry, overlap PCRs were performed to fuse a sentrin protease (SENP)cleavable N-terminal His6-SUMO tag, amplified from the pBAT4 derivative (45), pSAT1-LIC (this study), to either menT3 or menT4, amplified from H37Rv genomic DNA. The resulting PCR products were cloned as either Kpn I/Hind III fragments into Kpn I/Hind IIIdigested pBAD30 (menT3), producing pTRB517, or as Xma I/Hind III fragments into Xma I/Hind IIIdigested pBAD30 (menT4) to generate pTRB544.

Plasmids pPF656 and pPF657 were constructed by amplifying menA3 and menT3 from H37Rv genomic DNA and cloning as Mfe I/Xma I fragments into Eco RI/Xma Idigested pTA100 and pBAD30, respectively. To express His6-SUMO-tagged MenT3(D80A), site-directed mutagenesis was carried out using pTRB517 as a template. Briefly, nonoverlapping inverse primers were used to amplify menT3(D80A), followed by incubation with a mix of T4 DNA ligase, T4 polynucleotide kinase, and DpnI at 37C to remove template and circularize amplified DNA. This reaction was then used to transform E. coli DH5, resulting in pTRB593. Similarly, this method was used to generate MenT3(D80A), MenT3(K189A), and MenT3(D211A) for functional testing, using pPF657 as a template, resulting in pTRB591, pTRB562, and pTRB592, respectively.

Plasmid pTRB491 was generated by amplifying menA3 from H37Rv genomic DNA and cloning into pSAT1-LIC via ligation-independent cloning (LIC). The pSAT1-LIC plasmid features a LIC site that fuses an N-terminal His6-SUMO tag to the target protein. To produce MenT3(K189A) protein, the mutated gene was amplified from pTRB562 and similarly cloned into pTRB550 via LIC, resulting in pTRB577. The pTRB550 plasmid features a His6-SUMO LIC site, originally amplified from pSAT1-LIC and cloned as an Eco RI/Hind III fragment into Eco RI/Hind IIIdigested pBAD30.

To produce plasmids for use in M. tuberculosis, menA3, menT3, or both genes were amplified by PCR using PrimeSTAR GXL DNA polymerase, with M. tuberculosis H37Rv genomic DNA as template and primer pairs clo-RBS1-MenA3-attB2/clo-MenA3-attB3, clo-RBS1-MenT3-attB2/clo-MenT3-attB3, clo-RBS4-MenT3-attB2/clo-MenT3-attB3, or clo-RBS1-MenA3-attB2/clo-MenT3-attB3, respectively (tables S1 and S2). RBS1 (AGGAAGACAGGCTGCCC) and RBS4 (ACGAAGACAGGCTGCCC), corresponding to a strong or weak Shine-Dalgarno sequence, respectively, were placed upstream from the ATG translation start of MenA3 or the GTG translation start of MenT3. Plasmids pGMCS-TetR-P1-RBS1-MenA3, pGMCS-TetR-P1-RBS1-MenA3-MenT3, pGMCS-TetR-P1-RBS1-MenT3, or pGMCS-TetR-P1-RBS4-MenT3 were constructed by multisite gateway recombination (18), using plasmid pDE43-MCS as the destination vector. These plasmids are integrative vectors (insertion at the attL5 mycobacteriophage insertion site in the glyV tRNA gene) and express MenA3, MenT3, or MenA3-MenT3 under the control of P1 (Pmyc1 tetO), a tetracycline-inducible promoter (table S2) (46).

Construction of MenT3 D80A, D211A, and K189A substitutions for use in M. tuberculosis was performed as follows: Plasmid pGMCS-TetR-P1-RBS4-MenT3 was amplified by PCR with PrimeSTAR GXL DNA polymerase and the oligonucleotides pairs InFus-MenT3D80A-right/InFus-MenT3D80A-left, InFus-MenT3D211A-right/InFus-MenT3D211A-left, or InFus-MenT3K189A-right/InFus-MenT3K189A-left (table S1). The amplified linear fragments were purified on agarose gels and circularized using the In-Fusion HD Cloning Kit (Takara), as recommended by the manufacturer. Plasmids used to transform Stellar recipient cells were verified by sequencing and introduced by electroporation into M. tuberculosis (menA3-menT3)::dif6/pGMCZ (see the next paragraph).

Mutant strains of M. tuberculosis H37Rv were constructed by allelic exchange using recombineering (43), as previously described (fig. S2) (47). Two ~0.5-kb DNA fragments flanking the menA3-menT3 operon were amplified by PCR using PrimeSTAR GXL DNA polymerase (Takara), M. tuberculosis H37Rv genomic DNA, and the primer pairs MenA3Am-For/MenA3Zc-Am-Rev or MenT3Zc-Av-For/MenT3Av-Rev, respectively (table S1). A three-fragment PCR fused these two fragments to a Zc-resistance cassette flanked by two dif6 variants of the M. tuberculosis dif site and the recombination substrate was recovered by agarose gel purifications. The recipient strain for recombineering was a derivative of M. tuberculosis H37Rv carrying two plasmids: pJV53H, an Hm-resistant pJV53-derived plasmid expressing recombineering enzymes (43), and the integrative plasmid pGMCS-P1-MenA3, constitutively expressing menA3 (table S2). This strain was grown in complete 7H9 medium supplemented with Hm until mid-log phase and expression of recombineering enzymes was induced by Ace (0.2%) overnight at 37C. After induction, electrotransformation was performed with 100 ng of the linear DNA fragment for allelic exchange. After a 48-hour incubation at 37C, mycobacteria were plated onto agar supplemented with Zc. Zc-resistant clones were restreaked on the same medium, grown in complete 7H9 without antibiotic, and verified to be carrying the expected allele replacement by PCR amplification of chromosomal DNA and subsequent DNA sequencing, using primers MenA3Am-For/MenT3Av-Rev (fig. S1C and table S1). Spontaneous loss of the Zc-resistance cassette by XerCD-dependent recombination and of the pJV53H plasmid was obtained by serial rounds of culture without antibiotics and phenotypic tests for ZcS and HmS. Plasmid pGMCS-P1-MenA3 was then removed by transformation with pGMCZ, a similar integrative vector but carrying resistance to Zc, resulting in the deleted strain M. tuberculosis (menA3-menT3)::dif6/pGMCZ.

E. coli MC4100 dnaKdnaJ::KmR tig:CmR double mutant (40) was partially digested with Sau3 AI restriction enzyme and DNA fragments of about 1.5 to 4 kb in size were purified, then ligated into linearized and dephosphorylated Bam HIdigested pMPM2 (ColE1 origin) plasmid (39), and used to transform E. coli DH10B. About 25,000 independent transformants were pooled to constitute the multicopy library. This library has previously been used as a tool to identify multicopy suppressors of chaperone mutants (48).

In vivo toxicity and antitoxicity assays by cognate or noncognate antitoxins in E. coli were performed as follows. E. coli DLT1900 were cotransformed with pMPMK6-vector, pK6-MenT1, -MenT2, -MenT3, or -MenT4 (toxins), and p29SEN-vector, p29SEN-MenA1, -MenA2, -MenA3, or -MenA4 (antitoxins). Transformants were re-seeded from overnight cultures and grown at 37C to mid-log phase in LB supplemented with Km and Ap, and then serially diluted and spotted on LB-agar plates supplemented with Km and Ap, with or without l-ara (0.1%) and/or IPTG (200 M). Plates were incubated at 37C overnight and then imaged and counted. MenT3 substitutions were tested for toxicity in E. coli DH5 carrying pBAD30-vector, -MenT3 WT (pPF657), -MenT3(D80A) (pTRB591), -MenT3(K189A) (pTRB562), or -MenT3(D211A) (pTRB592). Strains were grown to mid-log phase, then serially diluted, and spotted onto M9M-agar plates supplemented with Ap, with or without l-ara (0.1%). After a 2-day incubation at 37C, plates were imaged and counted.

In vivo toxicity and rescue assays by cognate or noncognate antitoxins in M. smegmatis were performed as follows. Cultures of mc2 155 strain grown in LB at 37C were cotransformed with the integrative pGMC-vector, -MenT1, -MenT2, -MenT3, or -MenT4 (toxins), and with pLAM12-vector, pLAM-MenA1, -MenA2, -MenA3, or -MenA4 (antitoxins). Samples were selected on LB-agar plates supplemented with Km and Sm for 3 days at 37C, in the presence or absence of Atc (100 ng ml1) and Ace (0.2%) for toxin and antitoxin expression, respectively. A similar procedure was applied for pGMC-*MenA4-MenT4 carrying the menA4-menT4 operon, with the exception that no cotransformation with pLAM12 derivatives or selection on Km was needed.

Exponentially growing cultures [OD600 (optical density at 600 nm) between 0.05 and 0.2] of M. smegmatis strain mc2 155 containing plasmid pGMCS-TetR-P1-RBS1-MenT3 were divided in two: Half was left in complete 7H9 growth medium with Sm (uninduced cultures), while the other half was additionally treated with Atc (200 ng ml1) to induce expression from the P1 promoter. For labeling with LIVE/DEAD BacLight (Molecular Probes) dyes, cells were harvested 8 hours after Atc induction. Cells were centrifuged, resuspended in phosphate-buffered saline buffer, and stained as recommended by the manufacturer. Labeled cells were analyzed by fluorescence-activated cell sorting using a BD LSRFortessa X20 flow cytometer. Flow cytometry data analysis was performed using FlowJo software.

M. tuberculosis strains H37Rv or H37Rv (menA3-menT3)::dif6/pGMCZ were transformed by electroporation with 100 ng of plasmids pGMCS-TetR-P1-RBS1-MenA3, pGMCS-TetR-P1-RBS1-MenA3-MenT3, pGMCS-TetR-P1-RBS1-MenT3, pGMCS-TetR-P1-RBS4-MenT3, pGMCS-TetR-P1-RBS4-MenT3(D80A), pGMCS-TetR-P1-RBS4-MenT3(K189A), or pGMCS-TetR-P1-RBS4-MenT3(D211A). After 3 days of phenotypic expression in 7H9 ADC Tween at 37C, the transformation mix was divided into two halves. One half was plated on 7H11 OADC with Sm; the other half was plated on 7H11 OADC Sm supplemented with Atc (200 ng ml1). Plates were imaged after 20 days of incubation at 37C.

To perform in vivo copurification assays, E. coli BL21 slyD was transformed with (i) pET-MenT3-His, pET-MenA3-His, pET-MenT3/MenA3-His, or pET-MenT3-His/MenA3, or with (ii) pET-MenT1-His, pET-MenA1-His, pET-MenT1/MenA1-His, or pET-MenT1-His/MenA1, and selected on LB-agar plates supplemented with Ap and glu (20%). Transformants were grown at 37C to an OD600 of approximately 0.4 and then protein expression was induced overnight at 20C with 1 mM IPTG. Cell lysis and affinity purification of the protein complexes were performed as described below for MenT3-His purification. Elution fractions were separated on SDS-PAGE and proteins revealed using InstantBlue Protein Stain (Expedeon, catalog no. ISB1L).

To purify MenT3 for biochemistry, BL21 (DE3) slyD transformed with pET-MenT3-His, pET-MenT3-His(D80A), or pET-MenT3-His(K189A) was grown to an OD600 of approximately 0.4 at 37C. IPTG (1 mM) was then added, and the culture was incubated overnight at 20C. Under such conditions, MenT3 expression in E. coli was better tolerated and led to a reasonable amount of soluble MenT3 that could be collected for purification. Cultures were centrifuged at 5000g for 10 min at 4C, pellets were resuspended in Lysis buffer [300 mM NaCl, 50 mM tris (pH 7.5), and protease inhibitor tablet (Roche); 20 ml of buffer per 1 liter of cell culture] and incubated for 30 min on ice. Lysis was performed using the One Shot cell disrupter at 1.5 kbar (One Shot model, Constant Systems Ltd.). Lysates were centrifuged for 30 min at 30,000g in 4C, and the resulting supernatants were gently mixed at 4C for 30 min with Ninitrilotriacetic acid agarose beads (Qiagen, catalog no. 30230) preequilibrated with buffer PD [300 mM NaCl and 50 mM tris (pH 7.5)], using a 10-ml poly-prep column (Bio-Rad, catalog no. 7311550). Columns were stabilized for 10 min at 4C and washed three times with 10 ml of buffer PD plus 25 mM imidazole, and proteins were then eluted with buffer PD containing 250 mM imidazole. Elutions (500 l) were collected and PD MiniTrap G-25 columns (GE Healthcare, catalog no. 16924748) were used to exchange buffer with buffer PD supplemented with 10% glycerol. Proteins were concentrated using Vivaspin 6 columns with a 5000-Da cutoff (Sartorius, catalog no. 184501257). Proteins were stored at 80C until further use.

For additional MenT3 and MenT3(K189A) expression, either for crystallization or biochemistry, E. coli ER2566 pRARE pPF656 was transformed with either pTRB517 or pTRB577, respectively. For MenT3(D80A) expression, E. coli ER2566 pRARE was transformed with pTRB593. MenT4 was expressed in E. coli BL21 (DE3) transformed with pTRB544. MenA3 was expressed in E. coli ER2566 transformed with pTRB491. For these expressions, the same procedure was followed: Overnight cultures were re-seeded 1:100 into 2-liter flasks containing 1-liter 2 YT. Cells were grown at 175 rpm in 37C until an OD600 of 0.3 was reached and then at 22C until OD600 0.5, whereupon expression was induced by the addition of l-ara (0.1%) for toxins and IPTG (1 mM) for antitoxins. Cells were left to grow overnight at 16C, shaking at 175 rpm.

For selenomethionine incorporation, starter cultures of ER2566 pRARE pPF656 pTRB517 were grown overnight in LB at 37C with 200 rpm shaking. Cells were pelleted, washed, and resuspended in M9M, and then sub-cultured into 500 ml of M9M in 2-liter baffled flasks to a starting OD600 of 0.075. Cells were grown at 37C with 175 rpm shaking until an OD600 of 0.6, whereupon cells were centrifuged at 4200g and resuspended in fresh M9M. This sample was divided between separate 2-liter baffled flasks containing new M9M and shaken at 175 rpm for a further 1 hour at 37C. Once an OD600 of 0.7 was reached, 12 ml of nutrient mix [l-lysine hydrate (4 mg ml1), l-threonine (4 mg ml1), l-phenylalanine (4 mg ml1), l-leucine (2 mg ml1), l-isoleucine (2 mg ml1), l-valine (2 mg ml1), and 4 mM CaCl2] was added to each flask to promote feedback inhibition of methionine synthesis, followed by 250 SelenoMethionine Solution (Molecular Dimensions) to a final concentration of 40 g ml1, and cells were left to incubate for 1 hour at 20C. Last, toxin and antitoxin expression were induced by the addition of l-ara (0.1%) and IPTG (1 mM), and samples were left to grow overnight at 175 rpm in 16C.

All five proteins were purified in the same manner. Bacteria were harvested by centrifugation at 4200g, and the pellets were resuspended in buffer A500 [20 mM tris-HCl (pH 7.9), 500 mM NaCl, 5 mM imidazole, and 10% glycerol]. Cells were lysed by sonication at 40 kpsi and then centrifuged (45,000g, 4C). The clarified lysate was next passed over a HisTrap HP column (GE Healthcare), washed for 10 column volumes with A500, followed by 10 column volumes of buffer A100 [20 mM tris-HCl (pH 7.9), 100 mM NaCl, 5 mM imidazole, and 10% glycerol], and then eluted directly onto a HiTrap Q HP column (GE Healthcare) with buffer B100 [20 mM tris-HCl (pH 7.9), 100 mM NaCl, 250 mM imidazole, and 10% glycerol]. The Q HP column was transferred to an kta Pure (GE Healthcare), washed with 3 column volumes of A100, and then proteins were eluted using a gradient from 100% A100 to 100% buffer C1000 [20 mM tris-HCl (pH 7.9), 1000 mM NaCl, and 10% glycerol]. Fractions containing the protein peak were analyzed by SDS-PAGE, pooled, and incubated overnight at 4C with hSENP2 SUMO protease to cleave the His6-SUMO tag from the target protein. The following day, the samples were passed through a second HisTrap HP column and the flow-through fractions containing untagged target protein were collected. These samples were concentrated and run over a HiPrep 16/60 Sephacryl S-200 size exclusion column (GE Healthcare) in buffer S [50 mM tris-HCl (pH 7.9), 500 mM KCl, and 10% glycerol]. Peak fractions were analyzed by SDS-PAGE, pooled, and concentrated. Optimal fractions were separated and either flash-frozen in liquid N2 for storage at 80C or dialyzed overnight at 4C into buffer X [20 mM tris-HCl (pH 7.9), 150 mM NaCl, and 2.5 mM dithiothreitol (DTT)] for crystallographic studies. Crystallization samples were quantified and stored on ice and then either used immediately or flash-frozen in liquid N2 for storage at 80C. Frozen crystallization samples still formed usable crystals 15 months after storage.

Native and selenomethionine-derivatized MenT3 were concentrated to 12 mg ml1 and MenT4 was concentrated to 6 mg ml1, all in buffer X (see above). Initial crystallization screens were performed using a Mosquito Xtal3 robot (TTP Labtech) to set 200:100 nl and 100:100 nl protein:condition sitting drops. After initial screening and optimization, both MenT3 protein samples formed thick, six-sided needles in condition G5 [0.2 M calcium acetate hydrate, 0.1 M tris (pH 8.5), and 25% w/v polyethylene glycol 2000 monomethyl ether] of Clear Strategy II HT-96 (Molecular Dimensions). MenT4 formed thin, six-sided needles in the same condition as MenT3. To harvest, 20 l of condition reservoir was added to 20 l of cryo buffer [25 mM tris-HCl (pH 7.9), 187.5 mM NaCl, 3.125 mM DTT, and 80% glycerol] and mixed quickly by vortexing; an equal volume of this mixture was then added to the drop. After addition of cryo buffer, crystals were immediately extracted using a nylon loop and flash-frozen in liquid N2.

Diffraction data were collected at Diamond Light Source on beamlines I04 (MenT3 native), I03 (MenT3 selenomethionine-derivatized), and I24 (MenT4 native) (Table 1). Single 360 datasets were collected for native MenT3 and MenT4. Two 360 datasets from MenT3 selenomethionine-derivatized crystals measured at the selenium peak (0.9793 ) were merged using iSpyB (Diamond Light Source). Additional MenT3 selenomethionine-derivatized datasets were collected at selenium high remote (0.9641 ) and inflection (0.9795 ) wavelengths. Diffraction data were processed with XDS (49), and then AIMLESS from CCP4 (50) was used to corroborate the space groups (Table 1). The crystal structure of MenT3 was solved by MAD by providing the SHELX suite in CCP4 with the native and three anomalous MenT3 datasets. The solved starting model for MenT3 was built in REFMAC within CCP4. The crystal structure of MenT4 was solved ab initio using ARCIMBOLDO (51). Both models were then iteratively refined and built using PHENIX (52) and COOT (53), respectively. The quality of the final model was assessed using COOT and the wwPDB validation server. Structural figures were generated using PyMOL (Schrdinger). Comparison against models within the Protein Data Bank (PDB) was performed using DALI (25).

The following genetic procedure was developed and applied to select for E. coli genes that confer resistance to the MenT3 toxin. E. coli strain DLT1900 was first transformed with pK6-MenT3 (KmR) plasmid and transformants were selected at 37C on LB-agar plates supplemented with Km and glu (0.2%) to repress toxin expression from the araBAD promoter of pK6-MenT3. DLT1900 containing pK6-MenT3 was then grown in LB supplemented with Km and glu, transformed with the pMPMA2-based multicopy library of E. coli genes, and plated on selective LB-agar supplemented with Km, Ap, and l-ara (0.1%) to induce toxin expression. Plates were incubated for 24 hours at 37C. A control aliquot of transformants plated on nonselective plates (no l-ara) indicated that the number of transformants tested during the selection procedure was approximately 60,000. Note that under such conditions, E. coli DLT1900 pK6-MenT3 transformed with pMPMA2 empty vector did not produce any colonies on selective plates. We identified 72 toxin-resistant colonies that grew on selective plates after 24 hours, although they were smaller and translucent, indicating that growth inhibition by the toxin is not fully blocked by the suppressors identified. Of the 72 toxin-resistant colonies identified, only 41 were able to grow in culture. Plasmids were extracted from the 41 cultures, used to re-transform DLT1900 pK6-MenT3, and plated as above, to validate growth rescue in the presence of MenT3. Of 41 clones, 18 suppressors passed the second round of selection and were sequenced using the pMPMA2-For and -Rev primers (table S1).

Assays were performed as previously described (54). Briefly, template DNAs of DHFR (P0ABQ4), WaaF-Strep (P37692), and GatZ-Strep (P0C8J8) were used for in vitro transcription/translation coupled assays (PURExpress, New England Biolabs). These were performed according to the manufacturers instructions, in the presence or absence of the toxin. Following protein synthesis reactions of 2 hours at 37C, samples were separated on SDS-PAGE and visualized by InstantBlue staining (DHFR) or Western blots using anti-Strep tag antibodies (WaaF-Strep and GatZ-Strep).

Prevention of E. coli tRNATrp aminoacylation by MenT3 was monitored using a combination of two previously published methods (29, 55). E. coli BL21 (DE3) transformed with pETDuet or pET-MenT3 was grown at 37C to OD600 0.1 in M9M, whereupon expression of MenT3 was induced with 1 mM IPTG until an OD600 of about 0.4. The bacterial culture (25 ml) was then kept on ice and centrifuged for 10 min at 5000g in 4C. The pellet was resuspended in 0.5 ml of cold 0.3 M sodium acetate (pH 4.5) and 10 mM EDTA and transferred to a precooled 1.5-ml microcentrifuge tube, and 0.5 ml of phenol (equilibrated with the same buffer) was then added. After gentle pipetting, the sample was transferred into phase-lock tubes with an additional 400 l of cold chloroform. After 30 seconds shaking, the sample was first incubated on ice for 15 min and then centrifuged for 20 min at 20,000g in 4C. The aqueous phase was then transferred to a new cold 1.5-ml tube. Five hundred microliters of cold isopropanol was added and immediately mixed. RNA was precipitated for 1 hour at 20C, before the sample was centrifuged for 30 min at 20,000g in 4C (55). The supernatant was discarded and 1 ml of cold 75% ethanol was carefully added without disturbing the RNA pellet. After further centrifugation for 10 min at 20,000g in 4C, the supernatant was removed and the pellet was air-dried until no ethanol remained. The pellet was then resuspended by vigorously mixing in 20 l of cold 10 mM sodium acetate (pH 4.5) and 1 mM EDTA. Samples were stored at 80C. Samples were separated on a denaturing urea acrylamide gel for 3 hours at 100 V in 4C, as previously described (29). Northern blot and visualization with a radiolabeled DNA probe against tRNATrp was performed as previously described (56). Note that to distinguish the band of aminoacylated tRNA from its deacylated counterpart on the Northern blot, a chemically deacylated aliquot of RNA sample prepared from strain containing the empty vector was subjected to alkaline treatment. In this case, 46 l of tris-HCl (pH 9.0) was added to a 4-l aliquot of the RNA sample and incubated for 2 hours at 37C. Fifteen microliters of 0.3 M sodium acetate at pH 4.5 was added and followed by 125 l of 96% ethanol. RNA was precipitated at 20C for 1 hour, resuspended, and separated as described above.

For in vitro tRNA charging, in vitro transcription/translation assays were performed as above, using gatZ as DNA template. After a 2-hour reaction at 37C with or without MenT3 toxin (10 M), tRNA extraction, separation, and visualization were performed as described for the in vivo samples.

Labeled tRNAs were prepared by in vitro transcription of PCR templates containing an integrated T7 RNA polymerase promoter sequence. The template for E. coli tRNATrp was made by PCR amplification of chromosomal DNA from strain MG1655 with the primers CC2556 and CC2557 (CC2591 for tRNATrp without CCA) (table S1). The oligos for M. tuberculosis tRNAs are given in table S1. The T7 RNA polymerase in vitro transcription reactions were performed in 25-l total volume, with a 5-l nucleotide mix of 2.5 mM ATP, 2.5 mM CTP, 2.5 mM GTP, and 60 M UTP and 2 to 4 l of 10 mCi ml1 of radiolabeled UTP [-P32]. Template (0.1 to 0.2 g) was used per reaction with 1.5 l of rRNasin (40 U ml1) (Promega), 5 l of 5 optimized transcription buffer (Promega), 2 l of T7 RNA polymerase (20 U ml1), and 2.5 l of 100 mM DTT. Template DNA was removed by the addition of 2 l of RQ DNase (1 U ml1) (Promega). Unincorporated nucleotides were removed by G50 spin columns (GE Healthcare) according to the manufacturers instructions, in a final volume of 30 l. For E. coli tRNATrp, the transcript reaction was gel-purified on a denaturing 5% acrylamide gel and eluted in 0.3 M sodium acetate for 4 hours overnight at 4C. The supernatant was removed, ethanol-precipitated, and resuspended in 20 to 30 l of nuclease-free H2O.

MenT3 NTase activity was assayed in 10-l reaction volumes containing 50 mM tris-HCl (pH 9.5), 10 mM MgCl2, and 2.5 mM rNTPs and incubated for 20 min at 37C. Fresh, uniformly labeled tRNA (0.5 l) was used per assay, with different dilutions of the protein (1, 0.1, 0.01, and 0.001 mg ml1) in 50 mM tris-HCl (pH 7.8), 300 mM NaCl, and 10% glycerol. The 10-l reactions were mixed directly with 10 l of RNA loading dye (95% formamide, 1 mM EDTA, 0.025% SDS, xylene cyanol, and bromophenol blue), denatured at 90C, and applied to 5% polyacrylamide-urea gels. The gel was vacuum-dried at 80C and exposed to a PhosphorImager screen.

The effect of MenA3 antitoxin was assayed using in vitro-transcribed tRNASer2 as a substrate. For the coincubation assay, MenT3 (5 M) and increasing molar ratios of MenA3 were incubated with tRNASer2 and 2.5 mM CTP in 10-l reaction volumes containing 50 mM tris-HCl (pH 9.5) and 10 mM MgCl2 for 20 min at 37C. For the postincubation assay, the reactions were first incubated for 20 min at 37C with MenT3 alone in 7-l reaction volumes, then 3 l containing different concentrations of MenA3 were added, and the reactions were incubated for a further 20 min at 37C.

The tRNA screening was performed using 0.5 l of uniformly labeled M. tuberculosis tRNAs, all containing the CCA motif. The activity was tested in 50 mM tris-HCl (pH 9.5), 10 mM MgCl2, and 2.5 mM rCTP in 10-l reaction volumes and incubated for 20 min at 37C. The transcripts were incubated with 1 l of MenT3 (0.1 mg ml1), or with nuclease-free water as a control. The reaction was stopped with 10 l of RNA loading dye (95% formamide, 1 mM EDTA, 0.025% SDS, xylene cyanol, and bromophenol blue), denatured at 90C, and applied to 5% polyacrylamide-urea gels. The gel was vacuum-dried at 80C and exposed to a PhosphorImager screen.

Acknowledgments: We thank D.-J. Bigot for plasmid constructs, and P. Bordes, M.-P. Castani-Cornet, L. Falquet, L. Poljak, L. Hadjeras, and H. Akarsu for valuable advice. We also thank K. Semeijn and R. Dy for initial plasmid construction and testing, and E. Naser (Genotoul TRI-IPBS imaging facility) for help with flow cytometry analysis. Funding: This work was supported by a scholarship from the China Scholarship Council (CSC) as part of a joint international PhD program with Toulouse University Paul Sabatier (Y.C.); Springboard Award (SBF0021104) from the Academy of Medical Sciences (B.U. and T.R.B.); University of Otago Research Grant (P.C.F.), the School of Biomedical Sciences Bequest Fund, and University of Otago (P.C.F.); CNRS (UPR 9073), Universit Paris VII-Denis Diderot, the Agence Nationale de la Recherche (ARNr-QC), and the Labex (Dynamo) program (A.T. and C.C.); European Commission (contracts NEWTBVAC n241745 and TBVAC2020 n643381), Centre National de la Recherche Scientifique, Universit Paul Sabatier, Agence Nationale de la Recherche (ANR-13-BSV8-0010-01), and Fondation pour la Recherche Mdicale (DEQ20160334902) (C.G. and O.N.); and grant SNF CRSII3_160703 (P.G.). Author contributions: Conceptualization, all authors. Investigation, Y.C., B.U., C.G., A.T., and M. M. Writing, all authors. Funding acquisition, P.C.F., C.C., O.N., P.G., and T.R.B. Supervision, C.C., O.N., P.G., and T.R.B. Competing interests: The authors declare that they have no competing interests. Data and materials availability: The crystal structures of MenT3 and MenT4 have been deposited in the Protein Data Bank under accession numbers 6Y5U and 6Y56, respectively. All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors.

Go here to read the rest:
A nucleotidyltransferase toxin inhibits growth of Mycobacterium tuberculosis through inactivation of tRNA acceptor stems - Science Advances

INTRODUCTION

The important role of autophagy in health and disease has received unprecedented attention (1). As an essential and conservative physiological catabolic process, autophagy is responsible for the removal of protein aggregates, damaged organelles, and foreign bodies that invade cells (2). The autophagy contents are sequestered by double-membraned compartments (autophagosomes). Subsequently, the autophagosomes are fused with lysosomes to form autolysosomes, which degrade and circulate to produce nutrients (amino acids, fatty acids, and nucleotides) to be supplied to cells, and this dynamic process is called autophagic flux (3, 4). The unhindered autophagic flux is of great notable for maintaining homeostasis and protecting cells from attacks (5). The autophagic level is characterized by the amount of autophagy markers (e.g., autophagosomes and autolysosomes) and autophagy protein markers [e.g., LC3 (microtubule-associated protein-1 light chain-3)]. The upstream initiation or downstream blocking of autophagic flux will lead to the increase in autophagy markers (6). Malignant tumors are at a relatively high autophagic level compared with normal tissues to satisfy their metabolic demands, evasion, and resistance and allow tumor growth, survival, and malignancy (7). The involvement of autophagy in the occurrence and development of tumors suggests the reliable prospect of autophagy manipulation as an interventional means for tumor therapy (8). Autophagic flux blockade can disrupt the metabolism cycle of cancer cells, thereby reducing their fitness (9). Hydroxychloroquine (HCQ) and chloroquine (CQ) are the only clinically available autophagic blocking agents and they have been proven to be effective adjuvants for chemotherapeutics to increase their antitumor effects (10). HCQ/CQ, as a lysosomal alkalizing agent, can diffuse into lysosomes, causing the lysosomal pH to rise and dysfunction so that the lysosomes no longer fuse with the autophagosome, thereby blocking autophagic flux (11). However, the use of HCQ/CQ as a monotherapy strategy to block autophagic flux displays limited antitumor activity in clinical treatment (12). However, when HCQ is combined with autophagic stimulus, it can significantly increase its antitumor effect and reduce its dosage (13).

Aberrant endoplasmic reticulum (ER) status triggers autophagy stimulation (14). The ER acts as a reservoir of calcium ions in cells and is responsible for the correct folding and secretion of proteins (15). The accumulation and aggregation of misfolded proteins cause ER stress, triggering unfolded protein response (UPR) to significantly increase the autophagic level to restore homeostasis (16). As an ER stress initiator, tunicamycin (Tuni) blocks N-glycosylation and causes ER stressinduced autophagy to increase the autophagic level through PERK [protein kinase RNAlike ER kinase]/Akt (protein kinase B)/mTOR (mammalian target of rapamycin) signaling pathways, which are also closely related to matrix metalloproteinase-2 (MMP-2) expression (17).

Therefore, the combined application of the autophagic flux blocker HCQ and the ER stress initiator Tuni could cause cancer cells to have a special autophagic stress, which can severely disrupt cell homeostasis and cause cell death, resulting in a better therapeutic effect for tumor treatment. However, for a systemic administration of HCQ and Tuni, challenges such as poor hydrophilicity, poor biodistribution profile, low tumor accumulation, and tumor acid microenvironment prevent the drugs from penetrating the cell membranes, further affecting the application of free drugs in vivo (18). Moreover, high-dose HCQ in clinical application can only produce a moderate blockage of autophagic flux due to its poor efficacy, thus producing capricious therapeutic effects (19). Thus, the drug nanocarrier is considered to solve the above problems (20), as it has the following features: co-encapsulating hydrophilic and hydrophobic drugs, accumulating in tumor tissues through an enhanced permeability and retention (EPR) effect (21), entering the cells through the lysosomal pathway, and stimulating drug release with environment-responsive signals (22).

In addition, metastatic tumors undergo local migration and invasion in the early stage of tumor metastasis, and the migration speed depends on the focal adhesions (FAs) turnover (23). FAs are transmembrane multiprotein complexes containing integrins, paxillin, talin, zyxin, etc. Both cell-cell and cellextracellular matrix (ECM) adhesions form a stable connection via FAs (24). The decomposition of FAs at the cell rear by autophagy is critical for the forward movement and successful migration/invasion of tumor cells (25). In addition, MMPs are also an important factor in the selective regulation of tumor microenvironment to promote tumor metastasis, and are considered to be an inducer of epithelial-mesenchymal transition (26). The ER stress induced by Tuni regulates the PERK/Akt signaling pathway and down-regulates the expression of MMP-2 (27). Both the inhibition of the FAs turnover and the down-regulation of the expression of MMP-2 will reduce tumor metastasis.

Accordingly, in this study, we develop a pH-responsive polymersome for codelivering HCQ and Tuni drugs to simultaneously induce ER stress and block autophagic flux for achieving the antitumor effect and inhibiting tumor metastasis. A dual drugloaded, pH-responsive polymersome, Tuni/HCQ@CS-PAE, was designed to achieve this objective. The amphiphilic polymer chondroitin sulfate (CS)poly(-amino ester) was used to fabricate this polymersome, and hydrophilic HCQ and hydrophobic Tuni were loaded into the inner cavity and outer shell, respectively. Poly(-amino esters) with acid-stimulated responses are a class of highly biocompatible polymers and are believed to satisfy the performance of drug delivery (28). Repeated tertiary amine groups on the poly(-amino esters) are protonated by acid stimulation, thereby converting the hydrophobicity of the segment, resulting in the dissociation of the nanostructure and drug release (29). Simultaneously, the protonation process produces a similar HCQ effect to deacidify the lysosomes, swelling and rupturing the lysosomes, which can help the drugs to escape from the lysosomes and block the autophagic flux together with HCQ (30). Polymersomes with both a hydrophilic inner cavity and a hydrophobic shell are considered promising drug delivery platforms (31). In this work, the hydrophilic CS component can mask the surface positive charges of poly(-amino ester), prolonging the blood circulation time of the polymersomes. These polymersomes reached tumor tissues through the EPR effect, overcoming the nonselective distribution of the Tuni and HCQ drugs in vivo, increasing intratumoral accumulation, entering cells by endocytosis, and remaining in the lysosomes. Because of the pH response of poly(-amino ester), the polymersomes dissociated and produced a similar alkalization effect in lysosomes with HCQ, destroying and rapidly escaping the lysosomes, releasing the drugs Tuni and HCQ. Under the dual action of poly(-amino ester) and HCQ, the lysosomes in the tumor cells were destroyed, resulting in the blockade of autophagic flux. Moreover, the released Tuni triggered ER stress, further regulating the PERK/Akt signaling pathway to enhance the autophagic level and down-regulate the MMP-2 expression. The tumor cells were simultaneously attacked by both the inducement of autophagy due to the ER stress and the blockade of autophagic flux due to lysosomal destruction, resulting in a special autophagic stress, which seriously damaged the cell homeostasis and caused cell death.

The synthetic routes of CS-poly(-amino ester) are shown in fig. S1. The chemical structures of the block copolymers were confirmed using Fourier transform infrared spectra and nuclear magnetic resonance (NMR) spectra (figs. S2 to S4). The number-average molecular weight of CS-poly(-amino ester) was obtained as 8044 g/mol through gel permeation chromatography. The dual drugloaded Tuni/HCQ@CS-PAE polymersome was prepared by dialysis. The encapsulation efficiency (EE) and encapsulation content (EC) of Tuni were 38.5 and 10.5%, while those of HCQ were 56.1 and 11.2%, respectively. The morphologies of the Tuni/HCQ@CS-PAE polymersomes were characterized using transmission electron microscopy (TEM). Apparent vesicle structures with a size of ~180 nm can be observed in Fig. 1A, and the shell thickness of the vesicle in the enlarged image is ~30 nm. The average hydrodynamic size of the Tuni/HCQ@CS-PAE polymersomes was 230.0 9.3 nm, as measured using dynamic light scattering (DLS), with a polydispersity index of 0.156. The potential was 17.2 mV (Fig. 1B), and the negative charge indicated that it is suitable for drug carriers because it cannot be prematurely cleared in the blood circulation (32). The investigation of stability suggested that the polymersomes showed no significant changes (P > 0.05) in particle size when they were placed in phosphate-buffered saline (PBS) buffer and 10% fetal bovine serum (FBS) for 48 hours at 37C (fig. S5), indicating their potential for application in vivo. The pH response of the Tuni/HCQ@CS-PAE polymersomes was evaluated in a lysosomal acidic environment, and the CS-poly(-amino ester) was assayed to determine its pKa (where Ka is the acid dissociation constant) via acid-base titration. The results showed that CS-poly(-amino ester) had a pKa value of 5.4 (Fig. 1C), which was close to the lysosomal acidity (pH = ~5.0) (33). The TEM image obtained after the Tuni/HCQ@CS-PAE polymersomes were stored at pH 5.0 for 4 hours showed that the polymersome structure disappeared (Fig. 1D), indicating that the pH response of CS-poly(-amino ester) successfully caused the dissociation of the polymersomes. The 1H-NMR spectrum of CS-poly(-amino ester) in deuterium chloride at pH 5.0 (fig. S6) showed that the peak of poly(-amino ester) could be observed, indicating that it was hydrophilic at pH 5.0. DLS was further used to detect the pH response of the Tuni/HCQ@CS-PAE polymersomes. As shown in Fig. 1E, after the polymersomes were stored under the three acidic conditions of pH 7.4, pH 6.8, and pH 5.0 for 4 hours, the particle size distribution demonstrated that the polymersomes at both pH 7.4 and pH 6.8 (tumor ECM acidity) were stable; whereas at pH 5.0, the polymersome structure was destroyed, which is consistent with the TEM results. At pH 5.0, the potential of the Tuni/HCQ@CS-PAE polymersomes significantly shifted from 17.2 to 2.08 mV (Fig. 1F), indicating that the protonation of CS-poly(-amino ester) resulted in the capture of strong positive charges. The pH response of the polymersomes imparts them the ability to release drugs on-demand. As shown in Fig. 1 (G and H), the release of HCQ and Tuni at pH 6.8 was not significantly different (P > 0.05) from that at pH 7.4, suggesting that the polymersomes were stable in the tumor ECM and would not be released in advance. However, the 24-hour releases of HCQ and Tuni at pH 5.0 (lysosomal acidity) were 86.5 and 76.6%, respectively, which were 7.52 and 6.66 times the releases at pH 7.4, respectively. This result indicates that the Tuni/HCQ@CS-PAE polymersomes can rapidly release drugs in acidic lysosomes.

(A) TEM images of the Tuni/HCQ@CS-PAE polymersomes at pH 7.4. (B) Measurement results of the Tuni/HCQ@CS-PAE polymersomes by the Malvern laser particle size analyzer at pH 7.4. (C) Acid-base titration curve of CS-poly(-amino ester). (D) TEM images of Tuni/HCQ@CS-PAE at pH 5.0. (E) Hydrodynamic particle size distribution of the Tuni/HCQ@CS-PAE polymersomes at pH 7.4, pH 6.8, and pH 5.0. (F) potential of the Tuni/HCQ@CS-PAE polymersomes at pH 7.4, pH 6.8, and pH 5.0. (G) Release profiles of HCQ from the Tuni/HCQ@CS-PAE polymersomes. (H) Release profiles of Tuni from the Tuni/HCQ@CS-PAE polymersomes.

Before applying the polymersomes to cells and animals, both mouse breast cancer cells (4T1) and human umbilical vein endothelial cells (HUVECs) were used to evaluate the cytocompatibility of the polymersome delivery system. The blank material CS-PAE polymersomes exhibited good cytocompatibility at a concentration of 20 to 400 g/ml (cell viability over 85%, Alamar Blue assay; fig. S7, A and B). Only a small amount of red spots (representing dead cells) was observed in the fluorescence image of cells, with a polymersome concentration of up to 400 g/ml (live-dead cell staining; fig. S7C), also confirming the low cytotoxicity of the polymersomes.

The endocytic pathway of polymersomes was further examined in vitro. Fluorescein isothiocyanate (FITC)labeled (green) polymersomes were cocultured with adherent 4T1 cells, and the locations of the polymersomes in the cells and lysosomes labeled by LysoTracker Red DND-99 (red) were observed using fluorescence microscopy at 1 hour (fig. S8A) and 4 hours (fig. S8C), respectively. A large amount of yellow fluorescence in the cells was observed at 1 hour, which was the result of the overlap between green fluorescence and red fluorescence, suggesting that the polymersomes were in the lysosomes. At 4 hours, the yellow fluorescent signal decreased and the separated green and red fluorescent signals increased, indicating that the polymersomes were separated from the lysosomes. The figures (fig. S8, B and D) show the corresponding fluorescence intensity profiles of the white arrow regions in fig. S8 (A and C) obtained using ImagePro Plus, respectively. It can be observed that there was a large overlap between the two fluorescent signals at 1 hour, and their Pearsons correlation coefficient was calculated to be 0.88, indicating that the polymersomes and the lysosomes were strongly colocalized at 1 hour. At 4 hours, the Pearsons correlation coefficient was reduced to 0.04 according to fig. S8D, indicating that the polymersomes successfully escaped from the lysosomes. This result indicates that the polymersomes were endocytosed into the cells by the lysosomal pathway and could successfully escape the lysosomes at 4 hours in vitro.

The damage to the lysosomes by poly(-amino ester) and HCQ was marked by an increase in the lysosomal pH value. The LysoSensor Green-189 can monitor the acidity of the lysosomes, and its fluorescence reaches the highest value in normal lysosomes and decreases with increasing pH value. As shown in fig. S8E, it was observed via fluorescence microscopy that the green fluorescence in the HCQ, blank material CS-PAE polymersomes, and Tuni/HCQ@CS-PAE treatment groups was weakened to varying degrees. The results of the fluorescence-activated cell sorter (FACS) can be more intuitively observed (fig. S8F), and Tuni did not cause a change in the acidity of the lysosomes compared with that of untreated cells (control). Both the HCQ drug and CS-PAE polymersome had an alkalization ability for lysosomes, but HCQ performed better. This can be ascribed to the fact that CS-PAE polymersomes can only destroy lysosomes involved in endocytosis. The Tuni/HCQ@CS-PAE treatment group performed the best, and the median fluorescence intensity was only 14.3% of the control group, suggesting that the double action of HCQ and the polymersomes caused an increase in the intracellular lysosomal pH.

The relationship between various treatments and autophagy was further examined in vitro. Acridine orange (AO) is an acid-sensitive dye that stains the acidic organelles, including autophagosomes and autolysosomes in the cells red, whereas the DNA and cytoplasm in cells are green. Accordingly, the ratio of red to green signals can be used to evaluate the autophagic level (34). As shown in Fig. 2A, the number of red spots (observed via fluorescence microscopy) is positively correlated with the autophagic level. The red/green ratio calculated using FACS determines the autophagic level in each treatment group (Fig. 2B). The red/green ratios of the treatment groups increased compared with those of the untreated cells (control). The red/green ratio of the Tuni/HCQ@CS-PAE treatment group was approximately 68.0% higher than that of the Tuni/HCQ treatment group and was 1.91 and 2.21 times those of Tuni@CS-PAE and HCQ@CS-PAE, respectively. The red/green ratios of Tuni@CS-PAE and HCQ@CS-PAE also increased by 83.3 and 58.3%, respectively, compared with that of the blank material CS-PAE polymersomes. The red/green ratio of the CS-PAE polymersomes treatment group was also significantly increased by 71.4% compared with that of the control group. The results suggest that both Tuni and HCQ can cause an accumulation of acidic organelles, in addition to CS-PAE polymersomes. Unfortunately, the AO cannot distinguish whether the acidic organelles are autophagosomes or autolysosomes.

(A) Fluorescence images and (B) FACS analysis of AO-stained 4T1 cells after incubation with different treatments for 24 hours; ***P < 0.001. (C) Fluorescence images of mCherry-GFP-LC3 4T1 cells after incubation with different treatments for 48 hours. (D) Quantification of the number of LC3 puncta per cell (autophagosomes, yellow puncta; autolysosomes, red puncta). (E) TEM images of cells treated with saline or Tuni/HCQ@CS-PAE polymersomes (N, nucleus; green arrow, autophagosomes; red arrow, autolysosomes).

To track the autophagic flux, mCherrygreen fluorescent protein (GFP)LC3 adenovirustransfected cells were used. When autophagy occurs, a recognized autophagy marker, LC3, aggregates in both the inner and outer membranes of the autophagosomes. The LC3 in the transfected cells simultaneously expresses red fluorescence (mCherry) and green fluorescence (GFP). Thus, it will be observed in the form of yellow puncta in the autophagosomes. When the autophagosomes and lysosomes eventually fuse to form the autolysosomes, GFP is quenched by lysosomal acidity and only exhibits red puncta. Thus, the yellow and red puncta represent the autophagosomes and autolysosomes in the autophagic flux, respectively (Fig. 2C). The LC3 puncta statistic is shown in Fig. 2D. Comparing the LC3 puncta distribution of the Tuni@CS-PAE treatment group with that of the HCQ@CS-PAE treatment group, the former has more red puncta, indicating that the autophagic flux had entered the final stage, and the autolysosome had been formed. The large yellow puncta of the latter indicate that the autophagic flux mainly remained in the autophagosome stage, suggesting that HCQ@CS-PAE destroyed the lysosomes and prevented the autophagosomes from merging with the lysosomes. In contrast to CS-PAE polymersomes, Tuni@CS-PAE increased the autophagosome puncta and autolysosome puncta, indicating that it effectively increased the autophagic level in the cells. Compared with CS-PAE polymersomes, HCQ@CS-PAE showed a significant increase in the autophagosomes, but there was a decrease in the autolysosomes, which proved that HCQ@CS-PAE has a stronger ability to destroy the lysosomes and block the autophagic flux at the autophagosome stage. The autophagic level of the Tuni/HCQ@CS-PAE treatment group was greatly improved compared with that of Tuni/HCQ, whereas the number of autolysosomes was reduced. This result indicated that the dual drugloaded, pH-responsive polymersomes could increase the autophagic level and block the autophagic flux more evidently than free drugs.

From the TEM images (Fig. 2E), it can be observed that abundant autophagosomes (green arrows) and a few autolysosomes (red arrows) accumulated in the Tuni/HCQ@CS-PAE treatment group, in contrast to the control group (saline treatment). The increase in the autophagosomes and autolysosomes showed that the autophagic level of the Tuni/HCQ@CS-PAE treatment group was significantly enhanced. Furthermore, the amount of autophagosomes was significantly more than the amount of autolysosomes, indicating that the autophagic flux was blocked during the fusion process of the autophagosomes and the lysosomes.

The ability of the polymersomes to resist tumor metastasis was examined. A wound-healing assay (Fig. 3A) and the transwell invasion assay (Fig. 3B) were used to assess the cell migration and invasion in each group of treatments in vitro. The migration area (calculated by ImageJ) in the wound-healing assay is shown in Fig. 3C. The migration area of the Tuni/HCQ@CS-PAE treatment group is only 21.1% (P < 0.001) and 38.4% (P < 0.001) of those of Tuni@CS-PAE and HCQ@CS-PAE, respectively. Matrigel matrix (simulated ECM) was coated in the transwell upper chamber as an in vitro test tool for cell invasion. As shown in Fig. 3D, the number of cells (crystal violet staining) that arrived at the back of the polycarbonate membrane was counted as a quantitative index of invasive ability. The number of invasive cells in the Tuni/HCQ@CS-PAE treatment group was only 6.7% of that in the untreated group, 8.7% of that in the Tuni@CS-PAE treatment group, and 19.3% of that in the HCQ@CS-PAE treatment group. The above results show that the drug delivery system can effectively inhibit cell migration and invasion in vitro. Furthermore, Tuni@CS-PAE has a certain effect in inhibiting cell migration and invasion, but this effect is weaker than that of HCQ@CS-PAE.

(A) Typical images of wound-healing assay. (B) Cell invasion with the transwell assay (bottom). (C) Migration area of the wound-healing assay. (D) Number of invaded cells by the transwell assay. All data are represented as the means SD from three independent experiments; ***P < 0.001.

The in vitro cell viability of different treatment groups was firstly investigated. As shown in fig. S9A, the decrease in cell viability was positively correlated with increased autophagic level and autophagic flux blockade. On the basis of the concentration of fig. S9B, the median effect plots (fig. S9, C and D) of Tuni/HCQ and Tuni/HCQ@CS-PAE can be calculated to obtain their half maximal inhibitory concentration (IC50) values. According to the calculation method in the Supplementary Materials, the IC50 value of the Tuni/HCQ@CS-PAE treatment group is 8.2 M, which is 27.7% lower than that of the Tuni/HCQ treatment group.

The antitumor effect of the pH-responsive polymersome codelivering HCQ and Tuni drugs was further evaluated using the orthotopic luciferase genetransfected 4T1 (4T1-Luc) tumorbearing BALB/c mice. The treatment schedule is shown in Fig. 4A. After 7 days of orthotopic 4T1-Luc tumor implantation, the mice were randomly divided into six groups, and the drugs or drug-loaded polymersomes were administered intravenously at 0, 3, 6, and 9 days. The IVIS imaging system was used to monitor the bioluminescence signals of the tumors at days 0 and 30 (Fig. 4B). Ex vivo tumors were photographed on day 30 (Fig. 4C), and tumor volume was measured once in 3 days (Fig. 4D) and ex vivo tumors weighed on day 30 (Fig. 4E). Tumor growth inhibition (TGI; Fig. 4F) was calculated by tumor weight. The results indicated that three of the 4T1 tumors in the Tuni/HCQ@CS-PAE treatment group had successfully ablated, and the tumor weight was only 9.6% (P < 0.001) and 4.6% (P < 0.001) of those treated with Tuni@CS-PAE and HCQ@CS-PAE, respectively, whereas the TGI was as high as 97.5%. This proves that Tuni/HCQ@CS-PAE has an excellent antitumor effect. The body weight of the mice was monitored every 3 days (Fig. 4G), and it showed no difference in each treatment group on day 30 (P > 0.05), indicating the safety of the polymersome delivery system and the potential for application in vivo. Hematoxylin and eosin (H&E) staining of the main organs (heart, liver, spleen, and kidney) (fig. S10) also revealed no significant morphological changes in all the treatment groups.

(A) Treatment schedule for 4T1 breast tumor in BALB/c mice. (B) Bioluminescence images of 4T1-Luc tumorbearing BALB/c mice were taken on days 0 and 30 after various treatments. (C) Photographs of the tumors removed from the mice in the different treatment groups at the end of the experiment. (D) Tumor volume growth curves of the different treatment groups. (E) Weight of isolated tumors in the different treatment groups. (F) Tumor growth inhibition (TGI) after the different treatments. (G) Body weight changes of mice in the different treatment groups. (H) Hematoxylin and eosin (H&E) staining and immunohistochemical (IHC) analysis of tumor sections after the different treatments. All statistical data are presented as means SD. (n = 5; #P > 0.05; ***P < 0.001). [Photo credit for (B), (C), and (H): Funeng Xu, Southwest Jiaotong University].

H&E staining, terminal deoxynucleotidyl transferasemediated deoxyuridine triphosphate nick end labeling (TUNEL), and immunohistochemical (IHC) analyses (Ki67) were used to characterize the antitumor effects further (Fig. 4H). The H&E staining sections of the Tuni/HCQ@CS-PAE treatment group showed nuclear shrinkage and fragmentation, and the cell contour disappeared. Its TUNEL-positive signal (characterized apoptosis, brown) was the largest, and its Ki67-positive signal (characterized proliferation, brown) was the smallest. These results suggest that the lysosomal pH-responsive polymersomes entrapped with Tuni and HCQ can achieve excellent antitumor effects in tumor-bearing mouse models, demonstrating the success of autophagy regulation in antitumor applications.

The mouse 4T1 tumor is a metastatic tumor, corresponding mainly to lung and bone metastasis (35). The lung of the mouse was excised on day 30, and the lung metastasis of the tumor was observed using the bioluminescence images (Fig. 5A). There was no bioluminescence signal in the Tuni/HCQ@CS-PAE treatment group, indicating that there was no lung metastasis. Moreover, the pulmonary nodules are visualized using a Bouins fixative in Fig. 5B, which also supports this conclusion. The table summarizes the number of metastasis nodes (NOMN) (Fig. 5C), and the lung nodules are categorized by diameter: less than 0.5 mm, 0.5 to 1 mm, 1 to 2 mm, and greater than 2 mm, weighted 1 to 4 in turn. Because of the effective treatment of orthotopic tumors and the effective regulation of autophagy, Tuni/HCQ@CS-PAE has an excellent antimetastatic ability. The results of NOMN are shown in Fig. 5D. The average NOMN of the HCQ@CS-PAE treatment group was 52.89% that of the Tuni@CS-PAE treatment group and 22.49% that of the CS-PAE polymersomes treatment group. This indicates that both HCQ@CS-PAE and Tuni@CS-PAE can inhibit tumor metastasis to a certain extent, but the former performed better than the latter, which is also consistent with the results of in vitro antimetastasis evaluation. The H&E staining of the lungs can also demonstrate the antimetastatic effect (Fig. 5E). The red circle framed the foreign tissues of the lungs, and the foreign tissues were observed to be tumor tissues by comparison with the H&E staining of the tumors. No tumor tissue was observed in the H&E sections of the Tuni/HCQ@CS-PAE treatment group, and the area of the tumor tissue was significantly reduced in the Tuni@CS-PAE and HCQ@CS-PAE treatment groups compared with that in the saline group, which is consistent with the lung metastasis results of Fig. 5 (A and B).

(A) Bioluminescence images of tumor lung metastases in each treatment group in vitro. (B) Photographs of lung tissues; tumor metastasis was visualized by Bouins fixative, and metastatic nodules were white (represented by red arrows). (C and D) Counting the number of lung metastasis nodules, measurement of the diameter of metastatic tumors, and performing classification and counting. Number of metastasis nodes (NOMN) = I 1 + II 2 + III 3 + IV 4 (according to the diameter of the lung nodules for class 4: I < 0.5 mm, 0.5 mm II < 1 mm, 1 mm III 2 mm, and IV > 2 mm). (E) H&E staining of lung tissue after the various treatments. The red circle marks the metastatic tumor tissue. [Photo credit for (A), (B), and (E): Funeng Xu, Southwest Jiaotong University].

The dual drugloaded, pH-responsive polymersomes (Tuni/HCQ@CS-PAE) have made breakthroughs in the antitumor effect and metastasis inhibition effects in tumor-bearing mice. Western blot (WB) and IHC were further used to explore the mechanism of action of Tuni/HCQ@CS-PAE. ER stress and autophagy are closely related (Fig. 6A). It has been reported that the ER stress enhances the autophagic level by negatively regulating the Akt/mTOR pathway (36). In addition, the expression of MMP-2 is down-regulated by the down-regulation of Akt expression. Therefore, it can be concluded that the Tuni/HCQ@CS-PAE polymersome has two important functions in vivo: increasing autophagic levels and decreasing the MMP-2 expression and blocking the autophagic flow at the autophagosome stage by preventing the fusion of the autophagosomes and the lysosomes.

(A) Schematic diagram of Tuni causing ER stress, promoting autophagy, and reducing MMP-2 expression via signaling pathways in vivo. (B) WB analysis of key proteins of ER stress and downstream pathway protein of 4T1 tumor in BALB/c mice. (C) Relative expression level of key proteins in (B); ***P < 0.001 compared with control. (D) Expression of LC3 and p62 lanes of 4T1 tumor in BALB/c mice via WB. (E) Quantification of the ratio of LC3-II to glyceraldehyde-3-phosphate dehydrogenase (GAPDH) and p62 to GAPDH expression using ImageJ software. (F) IHC pictures of talin-1, paxillin, and MMP-2 in 4T1 tumorbearing BALB/c mice.

Under normal physiological conditions, GRP78/BiP (78-kDa glucose-regulated protein/immunoglobulin heavy chainbinding protein) acts as an ER chaperone and binds to ER receptors, which is in an inactive state. However, under ER stress, GRP78/BiP dissociates from ER receptors to activate and trigger UPR (37). The dissociation of GRP78/BiP from PERK (ER transmembrane receptor) triggers kinase dimerization and autophosphorylation to generate activated PERK (p-PERK) (38). Therefore, the expression of GRP78/BiP and p-PERK can be used as an indicator of ER stress. The expression of GRP78/BiP and p-PERK of the Tuni/HCQ@CS-PAE treatment group was significantly increased by 59.7% (P < 0.001) and 87.0% (P < 0.001), respectively, compared with that of the control group (Fig. 6, B and C). It indicated that Tuni/HCQ@CS-PAE can strongly trigger ER stress. The expressions of GRP78/BiP and p-PERK of the Tuni/HCQ treatment group were only 71.7% (P < 0.001) and 70.9% (P < 0.001), respectively, of that of the Tuni/HCQ@CS-PAE treatment group, suggesting that the polymersome system is more efficient than the free drugs at the same dose. The occurrence of ER stress triggered a series of downstream signaling pathways. The expressions of p-Akt and p-mTOR proteins in the Tuni/HCQ@CS-PAE treatment group were reduced by 80.0% (P < 0.001) and 67.2% (P < 0.001), respectively, compared to the control group, suggesting that the increase in the upstream event down-regulates Akt and mTOR activity. The expression of MMP-2 was significantly decreased in the Tuni/HCQ@CS-PAE and Tuni@CS-PAE treatment groups, which were only 32.8 and 41.0% of that of the control group, respectively, which was consistent with the trend of p-Akt expression.

When the autophagy occurs, LC3-I, which is in the cytoplasm of cells, is modified and processed to form LC3-II and expressed on the autophagosome membrane. The expression of p62 (sequestosome-1) as an autophagy substrate can characterize the smoothness of the autophagic flux. When the autophagic level is increased and the autophagic flux is smooth, the expression of LC3-II is increased and the expression of p62 is decreased; however, when the autophagy level is increased and the autophagic flux is blocked, the expression of both LC3-II and p62 is increased (39, 40). As shown in Fig. 6 (D and E), the expression of LC3-II and p62 was up-regulated in all the treatment groups compared with that in the control group, suggesting that the autophagic level of each treatment group was increased, and the autophagic flux was blocked to some extent. The highest expression of LC3-II was observed for Tuni/HCQ@CS-PAE and Tuni@CS-PAE, which was consistent with the results of down-regulation of p-mTOR. The expression of LC3-II of HCQ@CS-PAE was also significantly improved, indicating that, when autophagic flux was blocked by lysosomal destruction, the overall autophagic level of the cells also increased, and the autophagic flux mainly remained in the autophagosome stage. The expression of p62 was the highest in the Tuni/HCQ@CS-PAE and HCQ@CS-PAE treatment groups, i.e., 4.67 (P < 0.001) and 4.13 (P < 0.001) times that of the control group, respectively, suggesting that substrate degradation in the autophagosomes was largely blocked.

Talin-1 and paxillin are the constituent proteins of the FAs. The turnover of FAs is the basis of cell movement. The blockade of the autophagic flux leads to the failure of FAs degradation, which reduces the ability of cell movement. As shown in Fig. 6F, the IHC sections of Tuni/HCQ@CS-PAE and HCQ@CS-PAE showed the largest amounts of talin-1 and paxillin, indicating that the blockade of autophagic flux can significantly prevent FAs turnover. In the IHC section analysis of MMP-2, the expression of MMP-2 of the Tuni/HCQ@CS-PAE and Tuni@CS-PAE treatment groups was significantly inhibited, and the results were also consistent with the WB test in Fig. 6B, indicating that the MMP-2 expression was closely related to the ER stress induced by Tuni. It is suggested that based on the results of in vitro and in vivo antimetastasis experiments, it is known that HCQ@CS-PAE has stronger antimetastatic ability than Tuni@CS-PAE, also suggesting that the reduction in FAs turnover by autophagy affects tumor metastasis more than the decrease in the MMP-2 expression. Thus, Tuni/HCQ@CS-PAE achieved a considerable tumor-metastasis inhibition effect under the combined effect of down-regulation of MMP-2 expression and inhibition of FAs turnover.

CS was acetylated to increase its solubility in dimethyl sulfoxide (DMSO). Briefly, CS (1.0 g) was added to a round-bottom flask containing formamide (50.0 ml). The system was heated to 80C to promote dissolution and then cooled to room temperature. Pyridine (557.0 l) and acetic anhydride (500.0 l) were added to the round-bottom flask and magnetically stirred at room temperature for 12 hours. The reaction solution was dialyzed and lyophilized to obtain acetylated CS (Ac-CS; 0.91 g), which was stored for later use.

Ac-CS (0.41 g), 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide hydrochloride (0.16 g), and N-hydroxysuccinimide (0.04 g) were dissolved in reverse osmosis (RO) water. Triethylamine (0.08 g) was added to this mixture, and magnetic stirring was performed to activate the carboxyl group in an ice bath. Subsequently, 1,6-hexanediamine (0.63 g) was added, and the mixture was stirred at room temperature for 24 hours. The product Had-Ac-CS (0.49 g) was obtained via the dialysis and lyophilization of the reaction solution.

CS-poly(-amino ester) was prepared via the Michael addition reaction. Had-Ac-CS (0.37 g), Hexane-1,6-dioldiacrylate (HDDA) (0.5 g), and 3-dibutylamino-1-propylamine (DBPA) (0.37 g) were dissolved in 15 ml of DMSO, and the round-bottom flask was purged with N2. The mixture was stirred at 50C for 5 days. CS-poly(-amino ester) (1.1 g) was obtained via the dialysis and lyophilization of the reaction solution.

Dialysis was used to prepare the dual drugloaded polymersomes. First, 20.0 mg of CS-poly(-amino ester) and 7.0 mg of Tuni were codissolved in a beaker containing 5.0 ml of DMSO. Then, 5.0 ml of RO water containing 5.1 mg of HCQ was added dropwise under high-speed stirring, and the system was continuously stirred for 20 min. The system was transferred to a dialysis bag (molecular weight cutoff, 3500) for 48 hours and then lyophilized for the next experiment. EE and EC were calculated by Eqs. 1 and 2, respectivelyEE(%)=Weight of the drug in polymersomesWeight of the drug in feed100(1)EC(%)=Weight of the drug in polymersomesWeight of drug-loaded polymersomes100(2)

CS-poly(-amino ester) was placed in deionized water, and HCl solution was added until it was completely dissolved. Then, 1 to 5 l of 0.1 M NaOH solution was added dropwise, and the pH was measured after each addition. The pKa of the polymer is the pH at which it is half ionized.

The pH response of polymersomes was characterized by DLS, TEM, and 1H-NMR, respectively. The polymersomes were placed at pH 5.0, pH 6.8, and pH 7.4, respectively. After 4 hours, DLS was used to measure the particle size distribution and potential. Samples at pH 5.0 were also used for TEM and 1H-NMR detection.

The release of Tuni and HCQ in Tuni/HCQ@CS-PAE was investigated under three acidity conditions of pH 7.4, pH 6.8, and pH 5.0 at 37C. The release solution was taken 1 ml each time at the planned time point, and the same volume of fresh medium was added. The release solution was treated and analyzed by high-performance liquid chromatography.

Size changes of Tuni/HCQ@CS-PAE polymersomes after incubation in PBS or in cell culture medium [containing 10% FBS (v/v)] were monitored by DLS. The polymersome concentration was 1 mg ml1, and the experimental conditions were 37C.

Alamar Blue assay and live-dead staining were used to determine the cytocompatibility of blank polymersomes. In Alamar Blue assay detection, 1 104 cells per well of 4T1 and HUVECs were seeded in 48-well plates. After 24 hours of culture, polymersomes with 20 to 400 g/ml were added to each well. After 48 hours, the culture medium was removed, and 300 l of Alamar Blue solution [10% Alamar Blue, 80% media 199 (Gibco), and 10% FBS, (v/v)] was added for a further 3-hour incubation. They were then transferred to 96-well plates and detected by automated microplate spectrophotometer. For live-dead staining, 2 104 cells per well of 4T1 and HUVECs were seeded in 24-well plates. The cells were stained by 2 mM calcein acetoxymethylester for 10 min and propidium iodide for 5 min after 48 hours incubation, with different concentrations of polymersomes. Live cells were stained green, and dead cells were stained red when visualized by fluorescence microscopy.

4T1 cells were cultured in confocal dishes for 12 hours in Dulbeccos modified Eagles medium with 10% FBS. FITC-labeled polymersomes were added to two sets of confocal dishes, and then the medium was discarded at 1 and 4 hours, respectively. Cells were washed three times with PBS before staining, and then lysosomes were labeled with the LysoTracker Red DND-99. The fluorescence signal was observed by fluorescence microscopy.

Adherent 4T1 cells were treated with saline, Tuni, HCQ, CS-PAE, and Tuni/HCQ@CS-PAE for 12 hours, and stained with the acid-sensitive dye LysoSensor Green-189. The fluorescence intensity of each group was measured by fluorescence microscope and FACS.

The adherent 4T1 cells were treated with saline, CS-PAE, Tuni/HCQ, Tuni@CS-PAE, HCQ@CS-PAE, and Tuni/HCQ@CS-PAE, respectively. After 24 hours, the cells were stained with AO (1 l) for 15 min and detected by fluorescence microscopy and FACS, respectively.

4T1 cells were inoculated with 5 105 per well in confocal dishes before infection. The density of cells before virus transfection reached 50%, and the amount of virus mother liquor added to the plate was plaque-forming units = cell number multiplicity of infection (MOI). MOI was 20, 24 hours after infection; 2 ml of fresh medium was added to each well to replace the virus-containing medium.

4T1 cells were seeded in culture flasks at a density of 1 106 cells/ml for 18 hours. Then, saline and Tuni/HCQ@CS-PAE were added for 48 hours, respectively. Cells were digested, collected by centrifugation, and then fixed overnight in 2.5% glutaraldehyde. Samples were prepared according to TEM standard procedures and photographed.

Alamar Blue assay was used to assess the in vitro cytotoxicity. 4T1 cells were seeded in 48-well plates at a density of 2 104 cells per well. After the cells were cultured for 24 hours, various preparations were added to the well plates for 48 hours. After the incubation, 300 l of Alamar Blue solution was added for further 3 hours, and then the Alamar Blue solution was transferred to a 96-well plate, and the absorbance was measured with an automated microplate spectrophotometer. The median effect plot was a straight line fit with X = log(D) versus Y = log[fa/(1 fa)] (41). The theoretical IC50 value is the drug concentration corresponding to the x axis intercept of the median effect plot.

4T1 cells were seeded in six-well plates. When the cell confluence reached 100%, scratches were made with a 200-l pipette tip, and cells were washed three times with PBS to remove the delineated cells. The treatments were added to each group. Various therapeutic agents were added to the treated six-well plates and cultured for 36 hours in serum-free medium. The entire process was monitored with a microscope, and the healing area was calculated by ImageJ.

One hundred microliters of the diluted Matrigel was added vertically in the center of the transwell upper chamber and incubated at 37C for 4 hours to form a gel. Six hundred microliters of 10% serum medium was added to the lower chamber, and 100 l of the cell suspension was added to the upper chamber, and incubation was continued for 24 hours. The transwell chamber was removed, fixed in methanol for 30 min, stained with 0.1% crystal violet for 20 min, and the uninjured cells in the upper layer were gently wiped off with a cotton swab, and the count was observed with a microscope.

4T1-Luc cells were injected into the mouse mammary fat pad to establish an orthotopic breast cancer model. The animal experiments were approved by the Institutional Animal Care and Use Committee of Sichuan University and carried out in compliance with its guidelines. When the tumor volume of the mouse reached approximately 35 mm3, it was defined as 0 day of treatment, and the mice were randomly divided into six groups of five mice each. Each group was administered through the tail vein at 0, 3, 6, and 9 days of treatment [Tuni (7.5 mg/kg)]. Tumor volumes were calculated using the following equation: V = 0.5 A B2 (A refers to tumor length, and B refers to the tumor width). TGI was calculated using the following equation: TGI (%) = 100 (mean tumor weight of saline group mean tumor weight of experimental group)/mean tumor weight of saline group.

4T1 tumorbearing BALB/c mice were sacrificed, and the heart, liver, spleen, lung, kidney, and tumor tissues were excised, fixed with 10% formalin, dehydrated with gradient ethanol, and embedded in paraffin block. After denitrification with xylene, 4-m-thick tissue sections were stained with H&E, or for TUNEL detection, or for IHC staining with rabbit antiKi-67 polyclonal antibody and lastly observed with an optical microscope.

The potential association of ER stress with autophagy and antimetastatic mechanisms were analyzed by WB and immunohistochemistry. BIP/GRP78, PERK, Akt, mTOR, LC3, and p62 were used for WB analysis to explore the ER stressautophagy signaling pathway based on relative expression levels. MMP-2, talin, and paxillin were used as indicators of antimetastasis to analyze changes in their expression through WB and immunohistochemistry.

SPSS software was used for the statistical data analysis. Data were presented as means SD. One-way analysis of variance (ANOVA) was performed to determine statistical significance of the data. The differences were considered significant for #P > 0.05, *P < 0.05, **P < 0.01, and ***P < 0.001.

Acknowledgments: We thank the Analytical and Testing Center of the Southwest Jiaotong University. Funding: This work was partially supported by the China National Funds for Distinguished Young Scientists (51725303), the National Natural Science Foundation of China (21574105), and the Sichuan Province Youth Science and Technology Innovation Team (2016TD0026). Author contributions: F.X., Y.W., and S.Z. designed research; F.X., X.L., and X.H. performed research; F.X. and J.P. analyzed data; and F.X. and S.Z. wrote the manuscript. Competing interests: The authors declare that they have no competing interests. Data and materials availability: All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials. Additional data related to this paper may be requested from the authors.

Go here to read the rest:
Development of a pH-responsive polymersome inducing endoplasmic reticulum stress and autophagy blockade - Science Advances

INTRODUCTION

Photosynthesis transforms solar energy to chemical energy and supports nearly all life on Earth. Sunlight is absorbed by pigments in the light-harvesting (LH) antenna system, and excitation energy is transferred to the reaction center (RC), where photochemistry occurs, initiating an electron transfer process. The electron transport chain (ETC) couples the redox reactions associated with electron donors and acceptors to proton translocation to build up a proton motive force across the membrane, which, in turn, drives the formation of adenosine triphosphate (ATP) and other energy-consuming processes. In photosynthetic and respiratory ETCs, complex III (mitochondrial and bacterial cytochrome bc1, chloroplast and cyanobacterial cytochrome b6f) functions primarily to couple thermodynamically favorable electron transfer to proton translocation across the membrane (13). As quinol:electron acceptor oxidoreductases, these complexes create a transmembrane (TM) proton gradient through the Q-cycle mechanism: Four protons are translocated for every two electrons transferred to cytochrome c (cyt c) or plastocyanin upon quinol oxidation (35).

Notably, a functional counterpart for the cyt bc1 complex, alternative complex III (ACIII), has been identified in a wide range of bacterial taxa, and its presence usually coincides with the absence of the cyt bc1 complex (610). This complex is structurally and compositionally unrelated to the bacterial cyt bc1 complex, but it plays the same central role as a quinol:electron acceptor oxidoreductase in both the respiratory and photosynthetic ETCs (6, 813). In the respiratory chain, ACIII is usually associated with different cyt c oxidases and functions in aerobic electron transfer (14, 15). In the photosynthetic ETC of Chloroflexus aurantiacus, in which ACIII was originally discovered (16), the photosynthetic ACIII catalyzes the oxidation of menaquinol and mediates transfer of the released electrons to a periplasmic blue copper protein auracyanin, which, in turn, completes a cyclic electron transfer back to the RC (9, 12, 13).

Recent studies of the respiratory ACIII from Rhodothermus marinus (17) and Flavobacterium johnsoniae (18) have elucidated the structural features of this complex that are related to quinol coordination, cyt c oxidase association, and putative proton translocation. Regarding the association with different cyt c oxidases and the linear electron transfer mode of respiratory ACIII, the photosynthetic ACIII has a distinct composition and functions in a simple and efficient cyclic ETC using the electron donor menaquinol (8, 12, 13, 19, 20). However, the structure of the photosynthetic ACIII remains unknown. In particular, the fundamental coupling mechanisms underlying the menaquinol oxidation and proton translocation of the respiratory and photosynthetic ACIII complexes have received little research attention. Therefore, a structural investigation of the photosynthetic ACIII is necessary for a deeper understanding of the common coupling mechanism used by the ACIII from diverse bacterial taxa.

Roseiflexus castenholzii is a chlorosome-less filamentous anoxygenic phototroph. It contains a mosaic LH antenna, the type II pheophytin-quinone RC, and a cyclic electron transport system. The LH antenna of R. castenholzii is structurally similar to the LH1, but spectroscopically it resembles the LH2 of purple bacteria (19, 21, 22). Our previous structure of the R. castenholzii core complex (rcRC-LH) revealed a previously unknown architecture and a new type of menaquinone shuttling channel in the bacterial RC-LHs and illustrated the molecular basis underlying the LH and energy transfer mechanisms of early prokaryotes (23). We then extracted and purified R. castenholzii ACIII and its periplasmic electron acceptor auracyanin and revealed that ACIII oxidizes menaquinol-4 or menaquinol-7 and transfers the electrons to the copper ion coordinated in auracyanin (24). Here, we report the structures of the six-subunit R. castenholzii ACIII in air-oxidized and dithionite-reduced states, determined by single-particle cryoelectron microscopy (cryo-EM) at 3.3- and 3.5- resolution, respectively. We elucidated its structural features and here propose a previously unrecognized redox-coupled electron transfer and proton translocation mechanism that apparently links the respiratory and photosynthetic functions of the ACIII.

We purified the ACIII from phototrophically grown R. castenholzii using a modification of previous methods (7, 8, 16). We next used SDSpolyacrylamide gel electrophoresis (PAGE) and blue native PAGE to evaluate the purified ACIII (fig. S1A). Consistent with the molecular size expected from the corresponding gene sequences, we observed that the overall 300-kDa complex was composed of six subunits (ActA, ActB, ActC, ActD, ActE, and ActF), with molecular masses ranging from ~10 to 110 kDa (fig. S1A). Each subunit was confirmed by peptide mass fingerprinting (PMF) (tables S1 and S2). Further, gel filtration analysis (fig. S1B) indicated that the purified ACIII was a monomer containing one copy of each subunit. Spectral analysis indicated that the purified ACIII was air-oxidized: It was reduced via addition of sodium dithionite (fig. S1C). The reduced-minus-oxidized difference spectrum showed two absorbance peaks at 524 and 554 nm, indicating the increase of the c-type heme absorbance after reduction (fig. S1D).

To elucidate the proposed conformational changes that were involved in the redox-driven proton translocation of respiratory ACIII (17), the vitrified air-oxidized and dithionite-reduced ACIII were individually subjected to cryo-EM single-particle analysis. A dataset of 257,815 particles of the air-oxidized ACIII was used to reconstruct an electron potential map with an average resolution of 3.3 and a local resolution extending to 2.5 (fig. S2 and movie S1). The final reconstructed cryo-EM map was resolved and enabled us to build an accurate model of the TM helices with side chains in the air-oxidized ACIII (fig. S3) and all the cofactors and lipid molecules (Table 1 and fig. S4). The cryo-EM map of the dithionite-reduced ACIII was reconstructed from 488,581 particles to 3.5- resolution, the composition and overall structure of which resembles that of the air-oxidized ACIII (Table 1, fig. S5, and movie S2).

Different from the respiratory ACIII from R. marinus that contains seven subunits (ActABCDEFH) and one additional unidentified subunit (17), the photosynthetic ACIII only contains six core subunits (ActA, ActB, ActC, ActD, ActE, and ActF) as in F. johnsoniae (18). Superimposition of R. castenholzii ACIII structure with that of R. marinus and F. johnsoniae gives a main-chain root mean square deviation (RMSD) of 1.5 and 3.2 , respectively. Like the two respiratory ACIII structures, R. castenholzii ACIII is assembled into an L-shaped architecture with dimensions of 141 by 98 by 80 ; a TM arm (42 ) containing 23 TM helices from subunits ActA, ActC, ActD, and ActF; and a peripheral arm comprising the periplasmic subunits ActA, ActB, and ActE. On the periplasmic side, subunit ActB forms extensive contacts with ActE, the penta-heme binding domain of ActA, and the periplasmic region of ActC, ActD, and ActF. The loop between the two TMs of ActD inserts into the interface of ActA, ActB, ActC, and ActF. The globular domain of ActD is located in the cytoplasm (Fig. 1, A and B).

(A) The cryo-EM map of the air-oxidized ACIII is shown from the front (left) and back (right) view and represented with the dimensions of the TM and periplasmic region. Each of the six subunits is labeled with the length of its encoded amino acid. (B) Cartoon representation of the air-oxidized ACIII. The c-type hemes and lipids are shown as sticks, and the iron-sulfur clusters are shown as spheres. (C) Representation of the arrangement of cofactors in the air-oxidized ACIII. (D) Edge-to-edge distance between the iron-sulfur clusters and the hemes in the air-oxidized ACIII. The distances are labeled and shown in dashed lines. Color codes for all panels: lime green, ActA; slate, ActB; wheat, ActC; violet, ActD; yellow orange, ActE; aquamarine, ActF; red, c-type heme; orange brown, iron-sulfur clusters; yellow, lipids.

Similar to R. marinus ACIII, given its known electron transport function, one [3Fe-4S] and three [4Fe-4S] clusters in ActB, and six c-type hemes (five in ActA and one in ActE) were modeled in the density map of R. castenholzii ACIII (Fig. 1C and fig. S4), apparently forming wires of the six hemes and the iron-sulfur clusters (Fig. 1D). The six c-type hemes exemplify identical positions and orientations as that in R. marinus and F. johnsoniae ACIII, but there are only one [3Fe-4S] cluster and one [4Fe-4S] cluster identified at deviated positions in F. johnsoniae ACIII (fig. S6A). The six hemes and four iron-sulfur clusters are all located within reasonable edge-to-edge distances (less than 14 ) to permit direct electron transfer along the wire.

Structural superimposition of the air-oxidized and dithionite-reduced ACIII showed a main-chain RMSD of 0.4 (Fig. 2A), indicating that dithionite reduction does not induce obvious conformational changes at the current resolution. However, the difference map of ACIII (the air-oxidized map minus dithionite-reduced map) showed major electron potential differences at the periplasmic subunits (ActA, ActE, and ActB) and the cytoplasmic side of the TM region of ActA, ActC, and ActD (Fig. 2A). The increased electron potentials were observed at the six heme groups as well as the four iron-sulfur clusters (Fig. 2A and movie S3), indicating that these electron carriers are essentially reduced after dithionite treatment, which is consistent with the increased heme spectral differences upon dithionite reduction (fig. S1D).

(A) The air-oxidized minus dithionite-reduced electron potential difference map (orange) of the ACIII is shown from the front (left) and back (right) view. The structures of the air-oxidized and dithionite-reduced ACIII (white) are superimposed, with the iron-sulfur clusters and heme groups shown in sphere and stick models. The color code for each subunit and cofactors of the air-oxidized ACIII is the same as that in Fig. 1. (B) Ribbon representation of the ActA and ActE subunits bound with pentaheme and monoheme groups (red sticks). The N and C termini of the protein are highlighted with a black dot and labeled. (C) Spatial organization and immobilization of the pentaheme and monoheme groups in ActA and ActE subunits. The residues that axially coordinate the heme iron ions are shown as sticks and labeled; the center-to-center distances of the hemes are shown and labeled. (D) Overall structure of ActB subunit. The B1 and B2 domains are colored in blue and magenta, respectively. The iron-sulfur clusters are shown as spheres. (E) Coordination of the iron-sulfur clusters in the ActB subunit. The conserved cysteine residues that coordinate the iron-sulfur clusters are shown as sticks and labeled, and the B2 domain is shown as a ribbon with 80% transparency.

ActA (Gln9-Arg226) and ActE (Cys33-Asn193) were found to be penta-heme and mono-heme subunits, respectively, which form the main electron transfer wire of the photosynthetic ACIII. ActA is membrane-anchored, with an N-terminal TM helix (1, Gln9-Trp43). Five c-type hemes were bound in the loop regions between its six helices on the periplasmic side (Fig. 2B). The C-terminal mono-heme binding domain of ActE is composed of three helices and two turns (Fig. 2B), and our model showed a lipid anchor that is present at the N terminus of ActE (fig. S4D). This observation suggested the possibility that the consensus lipobox sequence L/V-A/T-G/A-C (M30TAC33) (fig. S6B) in the actE gene sequence may be excised from the transcript or degraded following translation in cells or at some point before the final complex assembly. This phenomenon was also observed in the respiratory ACIII (17).

The six heme groups bound by ActA and ActE are each covalently attached via thioester linkages to cysteine residues of highly conserved heme binding motifs (C-X-X-C-H), and their iron ions are axially coordinated through bi-His or His-Met residue couplets (Fig. 2C and fig. S6B). The five hemes of the ActA subunit are arranged in alternating parallel (heme_2,5 and heme_3,4) and perpendicular pairs (heme_2,3 and heme_4,5) (Fig. 2C). In particular, the heme_3,4 pair adopts typical stacked motif in van der Waals contact (edge-to-edge distance, 4.8 ), whereas heme_2,3 (5.1 ) and heme_4,5 (4.5 ) exemplify the T-shaped heme pairs (Figs. 1D and 2C). The spatial organization of heme_2 to heme_5 resembles that of the tetraheme in Shewanella oneidensis STC, in which the electron transfer between stacked heme pairs is approximately an order of magnitude greater than for the T-shaped heme pairs (25). But the electronic coupling of T-shaped heme pairs would be strongly enhanced by cysteine linkages inserted in the space between these pairs (26). The heme_1 is closest in terms of edge-to-edge distance to [3Fe-4S] (8.3 ), and it is buried in a hydrophobic pocket formed by residues from ActB, ActC, ActD, and ActE (fig. S6C). The porphyrin ring of the mono-heme in ActE is inclined about 60 compared to that of heme_5 in ActA, with an edge-to-edge distance of 9.0 and a center-to-center distance of 16.7 (Figs. 1D and 2C).

No midpoint redox potential data are available for the six hemes and iron sulfur clusters in R. castenholzii ACIII. The heme redox potentials of R. marinus ACIII was shown to range from 45 mV to +230 mV at neutral pH (11). Potentiometric titration of the c hemes in F. johnsoniae ACIII gives redox potentials at +331 mV and +439 mV (18). For C. aurantiacus ACIII, which shares 59% sequence identities with R. castenholzii ACIII, the heme midpoint redox potentials were determined to be 228 mV, 110 mV, +94 mV, and +391 mV (8). With the highest redox potential at +391 mV (8), the monoheme of ActE is believed to be the final electron prosthetic group to accept the electrons transferred from the five hemes in ActA. Regarding the high sequence homology and functional similarity (9) of ActA and ActE with that of C. aurantiacus and respiratory ACIIIs from R. marinus and F. johnsoniae (fig. S6, A to C), as well as the spatial distribution of the six hemes (Fig. 2, A to C), electrons can be sequentially transferred along a wire that begins with the heme_1 in ActA and ends with the monoheme in the ActE subunit, and then eventually to the acceptor auracyanin (13, 24).

The largest subunit, ActB (Gly77-Glu1006), was found to be composed of 26 helices and 17 strands that can be divided into two subdomains: the B1 domain (Gly77-Phe714) and B2 iron-sulfur binding domain (Leu715-Glu1006) (Fig. 2D). The N terminus of ActB was resolved from Gly77, just behind the signal peptidase cleavage site A71LA73. The twin-arginine translocase signal peptide assists with the translocation of ActB to periplasm (27). Superimposition analysis of ActB with PsrA and PsrB subunits of polysulfide reductase (PsrABC), an integral membrane-bound enzyme that performs quinone-coupled reduction of polysulfide substrates (28), revealed that the B2 iron-sulfur binding domain is similar to PsrB and that both the folding and positions of the four iron-sulfur clusters match well between the two subunits (fig. S7A). The analysis also revealed that the B1 domain of ActB forms a fold similar to the known substrate binding pocket of PsrA (fig. S9B), yet the absence of any cofactors in our model suggests that the function of ActB does not mirror the reduction activity of PsrA.

The four iron-sulfur clusters are covalently coordinated by conserved Cys residues (Fig. 2E and fig. S6D), with the largest edge-to-edge distance of 9.7 (Fig. 1D). The [3Fe-4S] is located at the interface with ActC and in the most proximity to the periplasmic side of the four-helix bundle that hosts the menaquinol binding pocket (Fig. 3A). This iron-sulfur cluster is the most probable primary electron acceptor from the menaquinol bound in the ActC subunit. The midpoint redox potential of [3Fe-4S] in R. marinus ACIII was determined to be +140 mV (11, 17), which is sufficient for an uphill electron transfer from menaquinol (70 mV at pH 7) (29). The role of the three [4Fe-4S] clusters in both respiratory and photosynthetic ACIIIs are still unknown. The air-oxidized minus dithionite-reduced electron potential differences at the [3Fe-4S] and three [4Fe-4S] clusters indicate that these iron-sulfur clusters can be reduced upon dithionite treatment (Fig. 2A and movie S3). An edge-to-edge distance of 8.3 was observed between the [3Fe-4S] and heme_1 in the ActC subunit (Fig. 1D), which suggests that the electrons accepted by the [3Fe-4S] cluster are most probably transferred along the heme wire to reduce a periplasmic electron carrier.

(A) Ribbon representation of the side (left) and bottom-up (right) views of the ActC (wheat) and ActF (aquamarine) subunits. The TM helices of ActC and ActF are labeled with numbers, and the iron-sulfur clusters in ActB and heme groups in ActA subunit are shown in spheres and red sticks, respectively. The N and C termini of each subunit are highlighted with a black dot and labeled. (B) Open cavity (bright yellow) between the TM helices of ActA, ActC, and ActD subunits of ACIII, which is equivalent to the menaquinol binding pocket. The cavity inside the TM region of ACIII was calculated using the program HOLLOW (44), and it is shown as a surface model. (C) Zoomed-in view of the putative menaquinol binding pocket, with essential amino acids shown as stick models. (D) Interactions between the modeled menaquinol head (blue stick model) and the menaquinol binding pocket. Residues are shown as stick models, and the hydrogen bonding interactions are shown as dashed lines with distances labeled.

The ActC (Lys8-Ala464) and ActF (Gln4-Ser399) subunits each contain 10 TM helices. The middle eight helices are arranged into two four-helix bundles (TM2-5 and TM6-9 of ActC, and TM2-5 and TM6-9 of ActF), which were sandwiched by the intersection of TM1 (TM1) and TM10 (TM10) (Fig. 3A). The helix bundles of ActC and ActF resemble the structure of PsrC dimer (fig. S7C). Superimposition of the structures of ActC and PsrC gives a main-chain RMSD of 1.1 . The quinone binding pocket of PsrC, which is formed by the N-terminal four-helix bundle and located at the periplasmic side, was identified according to the structures complexed with MK-7, pentachlorophenol, and ubiquinone-1 (28). Although no menaquinol was found in the current structures, we observed an open cavity between the TM helices of ActA, ActD, and TM3/4 of ActC subunits, which is equivalent to the quinol binding pocket of PsrC (Fig. 3B and movie S4).

On the basis of structural analysis and comparison as well as sequence alignment (fig. S8), we identified a menaquinol binding pocket of ACIII at the periplasmic side of the first four-helix bundle in ActC, about 12 away from the [3Fe-4S] cluster (Fig. 3C). Adjacent to [3Fe-4S], a strictly conserved His141 residue replaces the Glu67 of PsrC quinol binding pocket (Fig. 3C and figs. S7D and S8), which is involved in proton transfer from the menaquinol (30). The side chains of Trp84, Ile88, Phe91, Pro138, and Leu168 further form a hydrophobic pocket that is capable of immobilizing the menaquinol head group (Fig. 3D). The two carbonyl oxygen atoms of the modeled menaquinol head are capable of forming hydrogen bonds with the imidazole group of His141 (2.8 ) and the hydroxyl group of Asp171 (2.8 ), which further forms hydrogen bonds with Asp252 (2.9 ) (Fig. 3D). At the bottom of the pocket, Ile249 takes the position of Tyr130 in PsrC, which forms a hydrogen bond (2.6 ) with the O1 carbonyl group of MK-7 (fig. S7D) (28). The menaquinol binding pocket of R. castenholzii ACIII shares high sequence homology and conformational similarity with that of R. marinus and F. johnsoniae ACIII (fig. S7, E and F), indicating that ACIIIs play essentially similar enzymatic function in the photosynthesis and respiration.

On the basis of the structural comparison with the respiratory ACIII, we further identified a putative proton translocation passage in the ActC subunit. The passage begins at the cytoplasmic residues Arg198 and Asp199 and proceeds to the TM region located primarily at the first four-helix bundle of the periplasmic region (Fig. 4, A and C). This passage is composed of 22 proton-carrying residues that provide side chains for hydrogen bonding with protons (Fig. 4A). The air-oxidized minus dithionite-reduced electron potential differences were mainly distributed at the cytoplasmic side of TM1, TM3, TM4, TM5, and TM10 of ActC (Figs. 2A and 4C), where the menaquinol binding pocket and proton translocation passage are absent. Furthermore, we did not observe obvious structural differences at the proton translocation passage between the air-oxidized and dithionite-reduced structures (Fig. 2A).

(A) Organization of the menaquinol binding pocket (highlighted with a green box), [3Fe-4S] cluster, heme_1, and the putative proton translocation passage in ActC (wheat). The distance between [3Fe-4S] and the side chain of Asp171 in the menaquinol binding pocket is 12.2 , and the edge-to-edge distance between the [3Fe-4S] cluster and heme_1 is 8.3 . The residues that constitute the proton translocation passage and menaquinol binding pocket are shown as stick models, and the TM helices of ActC are shown as ribbon with 80% transparency. (B) Zoomed-in view of the hydrogen bonding networks between the menaquinol binding pocket and middle passage residues of the proton translocation passage in the ActC subunit, as well as the residues from the ActD and ActF subunits. Residues are shown as stick models, and the hydrogen bonding interactions are shown as dashed lines with distances labeled. (C) Topology diagram of the ActC subunit. The amino acids that constitute the menaquinol binding pocket and proton translocation pathway are shown in blue and green triangles, respectively.

In the middle of the passage, three strictly conserved residuesArg394, His246, and His99form a hydrogen bonding network that links the menaquinol binding pocket and proton translocation passage (Fig. 4B). The imidazole group of His246 forms hydrogen bonds with the guanidine group of Arg394 (3.1 ) and imidazole nitrogen of His99 (3.3 ), which forms a weak hydrogen bond with the main chain of Ile95 (3.3 ). The main-chain nitrogen of Ile95 is further hydrogen-bonded with the main-chain oxygen of Phe91 (3.1 ), one of the key residues involved in menaquinol coordination. In close proximity to His246, Ile248 forms a hydrogen bond with Asp171 (3.0 ), which is hydrogen-bonded with Asp252 (2.9 ) at the top of the menaquinol binding pocket (Fig. 4B).

Arg394, His246, and His99 are strictly conserved in both the respiratory and photosynthetic ACIII (fig. S8). Superimposition analyses showed that the triplet residues adopt the same side-chain orientations and hydrogen bonding network as that from R. marinus and F. johnsoniae (Fig. 5, A and B), suggesting that these residues share a similar function in the respiratory and photosynthetic ACIII. Asp394 is also conserved in other polysulfide, tetrathionate, nitrate, and dimethyl sulfoxide reductases (30). Mutation of Arg394 in Wolinella succinogenes PsrC resulted in an inactive enzyme, which was suggested that it stabilizes the deprotonated quinol (30). Regarding the sequence conservation, location, and extensive hydrogen bonding interactions with the menaquinol binding pocket, the triplet residues are likely essential for coupling the menaquinol oxidation and proton translocation.

(A) Comparison of the proton translocation passage in R. castenholzii ActC (wheat) with that of the respiratory ActC from R. marinus (PDB 6f0k, white) and F. johnsoniae (PDB 6btm, pink). The [3Fe-4S] cluster, heme_1, and the amino acids are shown as sticks. (B) Middle passage residues that are capable of forming hydrogen bonding networks. (C) Putative proton translocation passage in R. castenholzii ActF subunit (aquamarine) and its superimposition with that of R. marinus (PDB 6f0k, white) and F. johnsoniae (PDB 6btm, pink). (D) The middle passage residues that are capable of forming hydrogen bonds in ActF subunits are shown in stick models.

We observed a similar proton translocation passage in the ActF subunit formed by 20 less conserved amino acids (about 20% identities) from the cytoplasmic to periplasmic side (Fig. 5C and fig. S9). In the middle of the ActF passage, side chains of Glu335, Ser217, and Tyr339 are capable of forming hydrogen bonding interactions, but no menaquinol-binding pocket and similar hydrogen bonding networks as that in ActC were found (Fig. 5C). In addition, Ser217 and Tyr339 are less conserved in both the photosynthetic and respiratory ActF, and Glu335 is replaced by Arg in C. aurantiacus and His residue in the respiratory ActF (Fig. 5D and fig. S9). Minor electron potential differences were only observed at His287, Ala189, and Met65 of the ActF subunit, suggesting that this subunit is not sensitive to the air-oxidized and dithionite-reduced state of ACIII. To be noted, a conserved residue Tyr264F forms a hydrogen bond with the main-chain oxygen of Pro267C (2.8 ), which is close to the periplasmic portion of the proton translocation passage in ActC (Fig. 4B). The distinct conservation of these proton translocation passages indicates that the ActC subunit plays consensus important role in both the respiratory and photosynthetic ACIII.

The function of the ActD subunit in the ACIII remains obscure. We observed hydrogen bonding interactions between Asn100 of ActD and Tyr755 of ActB, as well as between Leu106 of ActD and Tyr753 of ActB (fig. S10A). We also observed extensive hydrophobic interactions between residues located in the ActD loop and subunits ActB, ActF, and ActC. These interactions can stabilize the conformation of TM5, which contributes to the menaquinol binding pocket (fig. S10A). Near the menaquinol binding pocket, a hydrogen bond is formed between the hydroxyl groups of Glu118 of ActD and Ser244 of ActC (2.1 ), which was close to the His246 of ActC that would be essential for coupling the menaquinol oxidation and proton translocation (Fig. 4B). Thus, ActD might play a primary role in stabilizing the TM region of ACIII, which thereby contributes to a stable menaquinol binding pocket and proton translocation passage.

As a functional counterpart of the bc1 complex, ACIII plays a central role in both the photosynthetic and respiratory ETC of a wide range of bacterial taxa (610). It couples quinol oxidation with TM proton translocation to build up a TM proton gradient, which drives the formation of ATP required for bacterial growth. However, the nature of the coupling mechanism(s) for the respiratory and photosynthetic functions of ACIII has not been well discussed.

The photosynthetic bacterium R. castenholzii has evolved a simple but efficient cyclic ETC to transform solar energy into chemical energy that is different from the linear respiratory chain (3133). Our study has revealed the structure of the first photosynthetic ACIII comprising six conserved subunits, in both the air-oxidized and dithionite-reduced states, as well as the nature and position of the cofactors, including six hemes and four iron-sulfur clusters. We also detected a menaquinol binding pocket positioned at the periplasmic side of the TM subunit ActC. This pocket is capable of immobilizing the menaquinol head group via strictly conserved residues (Fig. 3D), which is linked by extensive hydrogen bonding interactions with three proton-carrying residues in the middle of an apparent proton translocation passage. In addition, the ActD subunit is shown to coordinate extensive interactions with subunits ActA, ActB, ActC, and ActF.

Previous enzymatic analyses confirmed the activity of photosynthetic ACIII as a menaquinol:auracyanin or cyt c oxidoreductase (9). Recently, we revealed that R. castenholzii ACIII oxidizes menaquinol-4 or menaquinol-7 and transfers electrons to its periplasmic electron acceptor auracyanin (24). It has been revealed that there is a single quinol binding site in R. marinus ACIII by isothermal titration calorimetry experiments (17). The high sequence and structural similarity among photosynthetic and respiratory ACIIIs would also suggest a single menaquinol binding pocket of R. castenholzii ACIII. Within this pocket, menaquinol binds and is oxidized by the terminal electron acceptor auracyanin, releasing two protons into periplasm. Considering that menaquinone is reduced at the binding site of RC-LH complex (23), accepting two protons from cytoplasm, an apparent efficient quinone shuttling cycle is formed among RC-LH, the membrane quinone pool, and ACIII in the R. castenholzii simple cyclic photosynthetic ETC. As a result, with the reduction of one molecule menaquinone at RC-LH and the oxidation of one shuttled menaquinol at ACIII, two transferred electrons are accompanied with two protons transferred from cytoplasm to periplasm, yielding a H+/e ratio of 2:2.

To date, no experimental data on the H+/e stoichiometry for any ACIII were reported. Previous studies proposed that ACIII could also actively pump additional protons from cytoplasm into periplasm (10, 12, 14, 15), which would yield a different H+/e stoichiometry deduced from above quinone shuttling cycle. However, the detailed mechanism of its active proton translocation has not been elucidated. The lack of any redox-active cofactors in the TM and cytoplasmic regions of ACIII argues against a Q-cycle type H+ pumping mechanism, such as is used in the cyt bc1 and cyt b6f complexes.

Structural comparison and analyses revealed two putative proton translocation passages in ActC and ActF, respectively, for both photosynthetic and respiratory ACIII (Fig. 5, A and C). The side chains of the middle-passage triplet residues Arg394, His264, and His99 of ActC adopt exactly the same conformation for all three ACIIIs (Fig. 5B). However, the proton-carrying residues in the passage of ActF are less conserved than that of ActC (Fig. 5, C and D). Notably, the respiratory ACIIIs from R. marinus (17) and F. johnsoniae (18) contain two conserved His and Asp residues in the middle passage of ActF, but these two residues are replaced by Glu and Tyr in the R. castenholzii ACIII (Fig. 5D and fig. S9). In addition, neither menaquinol binding pocket nor hydrogen bonding network was found in ActF. Less differences of electron potential around ActF between the air-oxidized and dithionite-reduced states (Fig. 2A) suggest that ActF is insensitive to the changes of redox potential. Therefore, most probably, ActF passage lacks a driving force for efficient TM proton translocation. If there exists a redox-coupled active proton translocation in ACIII, it would be mostly located in the ActC subunit and driven by the coupling between menaquinol oxidation and putative proton passage, without the necessary conformational change.

On the basis of the above structural analysis and discussion, we propose a redox-coupled proton translocation mechanism for the photosynthetic ACIII, which occurs within the subunit of ActC (Fig. 6). In the menaquinol binding pocket, at the close-to-neutral pH environment (pH ~6.5) of periplasmic space, both Asp171 and His141 are deprotonated and coordinate the bound menaquinol (MQH2) by hydrogen bonds. The hydroxyl hydrogens of menaquinol can be bound by the hydroxyl oxygen of Asp171 and imidazole nitrogen of His141, respectively. Upon oxidation, the hydroxyl group of menaquinol that faces the side chain of Asp171 is first oxidized to form an intermediate semi-menaquinol. The released hydrogen protonates Asp171. Lacking the coordination by Asp171, the semi-menaquinol would be relocated in the binding pocket and thus enable extraction of one proton from the proximal proton passage of ActC, resulting in one proton translocated from the cytoplasm. The binding of the extracted proton will induce a reorganized electronic structure of semi-menaquinol, releasing another hydroxyl hydrogen to protonate His141. The reorganized semi-menaquinol can be further coordinated by the hydroxyl group of Asp171. Then, the semi-menaquinol is further oxidized to form menaquinone (MQ) and release the exacted proton. After the release of menaquinone and the extracted proton from the menaquinol binding pocket, the two protons from oxidation of menaquinol are released to periplasmic space with the deprotonation of Asp171 and His141. During this proposed process, one instance of menaquinol oxidation is coupled to one proton pumped from the cytoplasm. As a result, three protons are released into the periplasm per two electrons transferred (Fig. 6).

The menaquinol head group and the side chains of the essential amino acids are shown to indicate the coupling mechanism of the photosynthetic ACIII from R. castenholzii. Upon menaquinol oxidation, two electrons are sequentially transferred to the [3Fe-4S] cluster with a time interval, two protons from oxidation of menaquinol are released to periplasmic space, and one proton is pumped from the cytoplasm (colored in red) through hydrogen bonding networking with the essential amino acids in the proton translocation passage. As a result, three protons are released into the periplasm per two electrons transferred during oxidation of one instance of menaquinol.

In both the respiratory and photosynthetic ACIII structures, a [3Fe-4S] cluster in the ActB subunit functions as the primary electron acceptor from menaquinol (17, 18), donating the electrons along the six-heme wire and finally onto the periplasmic electron acceptor. Both the photosynthetic ACIII from R. castenholzii and the respiratory ACIII from R. marinus contain additional three [4Fe-4S] clusters, while only one [4Fe-4S] cluster was identified in the F. johnsoniae ACIII (18). The function of [4Fe-4S] clusters remains largely unknown.

Our observation of the electron potential differences of these [4Fe-4S] clusters between air-oxidized and dithionite-reduced states indicates that these clusters are either accessible to dithionite or connected to the electron transfer wire. In Psr with the absence of heme groups, two electrons released from MK-7 are transferred via five [4Fe-4S] clusters to the bis-MGD (bis-molybdopterin guanine dinucleotide) cofactor and then reduce polysulfide (28). Unfortunately, no cofactors were observed in the B1 domain of ActB subunit (fig. S7B), indicating an electron transfer dead end in these [4Fe-4S] clusters. How they contribute to the electron transfer of ACIII needs to be further considered.

Both heme and iron-sulfur cluster are single electron carriers that are unable to transfer two electrons simultaneously. Thus, a sequential transfer of electrons upon menaquinol oxidation is necessary. In addition, the latency time between the formation of semi-menaquinol and its further oxidation needs long enough to allow extraction of proton from the translocation passage, but it should not be too long to avoid the formation of reactive oxygen species. On the other side, the final periplasmic electron acceptor auracyanin can only accept one electron each time. Therefore, the speed of electron transfer in ACIII should be well controlled. The alternating T-shaped spatial organization of the six hemes in ACIII would limit in one order the electron transfer efficiency of the heme wire, which would increase the steady time of semi-menaquinol. This limitation could be further compensated by the [4Fe-4S] clusters playing as an electron sink. Overall, the possible electron transfer during menaquinol oxidation would look like that, the first electron would quickly sink into the [4Fe-4S] clusters via [3Fe-4S] with the formation of semi-menaquinol, and the second electron could then be transferred to the final periplasmic acceptor auracyanin via the heme wire; with a second auracyanin binding, the sinking electron in the [4Fe-4S] clusters could be further transferred to the final acceptor via the heme wire. As a result, the existence of the [4Fe-4S] clusters would be very important in assisting sequential and efficient transfer of two electrons with an intrinsic time interval.

In summary, our work provides a structural basis and conceptual insight into the coupling mechanism underlying menaquinol oxidation, electron transfer, and proton translocation for the photosynthetic ACIII, which seems likely to play the same role as a menaquinol:electron acceptor oxidoreductase in respiratory ACIIIs. Direct experimental will be required for definitive characterization the proton pumping mechanism of these ACIIIs.

R. castenholzii DSM 13941 was grown in a batch culture anaerobically in modified PE medium at 50C under high-light conditions for 10 days (19). Cells were harvested by centrifugation at 10,000g for 20 min, and the pellet was washed twice with 20 mM tris buffer (pH 7.4) and then stored at 40C.

A suspension of whole membranes [with OD880 (optical density at 880 nm) = 20 cm1] in 20 mM tris-HCl (pH 8.0; buffer A) was treated with 1% -octyl glucoside and stirred for 1 hour at room temperature in the dark. The extraction was centrifuged at 200,000g for 2 hours (Ti 70 rotor, 45,000 rpm) at 4C. The pellets were resuspended in 50 mM sodium acetate (pH 5.0; buffer B) and treated with 0.5% -dodecyl maltoside as above with 1% -octyl glucoside. The supernatant from the second ultracentrifugation was collected and filtered through a 0.22-m Millipore filter and subsequently loaded on a prepacked cation exchange chromatography column (SPHP5, GE Healthcare), which had been equilibrated with buffer B containing 0.04% -dodecyl maltoside (which makes up buffer C). The column was extensively washed with 50 mM NaCl in buffer C until the eluent was colorless. Last, the crude ACIII was eluted from the column by a sodium gradient from 0.1 M NaCl to 0.4 M NaCl with 50 ml of buffer C at 2 ml min1. The collected fractions were concentrated and further purified by Superdex-200 gel filtration in buffer D [100 mM NaCl, 0.02% -dodecyl maltoside, and 20 mM tris-HCl (pH 8.0)]. The fractions with an absorption ratio of A413/A280 higher than 1.38 were pooled and used for cryo-EM analysis.

The polypeptide composition of the purified complex was determined by SDS-PAGE and blue-native PAGE. The sample solubility was optimized by dissolving samples in buffer containing 5% 2-mercaptoethanol for 30 min at 65C; these conditions yielded the sharpest protein bands. The identity of SDS-PAGE and blue-native PAGE bands was confirmed by PMF using matrix-assisted laser desorption/ionizationtime-of-flight (MALDI-TOF) mass spectroscopy.

Stained bands from the SDS-PAGE were excised and destained and washed with 50% acetonitrile in 50 mM aqueous NH4HCO3. Proteins were then reduced with 10 mM dithiothreitol in 100 mM NH4HCO3 for 30 min. Cysteine residues in the proteins were further alkylated by 55 mM iodoacetamide in 100 mM NH4HCO3 for an additional 30 min. Trypsin (Promega Trypsin Gold, TPCK (L-1-tosylamido-2-phenylethyl chloromethyl ketone)treated) in 50 mM NH4HCO3 was added to the gel pieces, and the enzymatic reaction proceeded overnight at 37C. Afterward, peptides were extracted twice with 1% trifluoroacetic acid in 60% acetonitrile for 30 min. Extracted solutions were collected, dried completely in a speed-vac, and then redissolved in 50% acetonitrile containing 0.1% trifluoroacetic acid for mass spectrometry analysis.

The identities of proteins were determined by PMF using an ABI 4700 MALDI-TOF mass spectrometer. A mixture of the peptide sample and freshly prepared matrix solution (10 mg ml1 -cyano-4-hydroxycinnamic acid in 50% acetonitrile) was spotted on a stainless-steel target plate. Peptide mass value searches were performed against the National Center for Biotechnology Information (NCBI) database using Mascot software (www.matrixscience.com). The alkylation of cysteine was included as a possible modification. The mass tolerance for the monoisotopic peptide mass was set to 0.6 Da.

Three-microliter aliquots of air-oxidized ACIII (4 mg ml1) was placed on the glow-discharged GiG R1.2/1.3 300-mesh gold holey carbon grid (Jiangsu Lantuo Biotechnology, China) and blotted for 3.0 s under a blot force of 1 at 100% humidity and 16C before being flash-frozen in liquid ethane with a Mark IV Vitrobot system (FEI). Micrographs were acquired on a Titan Krios microscope (FEI) operated at 300 kV with a K2 Summit direct electron detector (Gatan). SerialEM (34) was used for automatic data collection. A nominal magnification of 22,500 was used for imaging, which yielded a pixel size of 1.04 . The defocus range was between 1.2 and 3.3 m. Each micrograph was dose-fractionated to 32 frames under a dose rate of 9.2 e/2 per second and an exposure time of 6.4 s, which resulted in a total dose of about 59 e/2.

For the sodium dithionitereduced ACIII, 3-l aliquots of a sample (4.5 mg ml1) were placed on the glow-discharged CryoMatrix R1.2/1.3 300-mesh amorphous alloy film (product no. M024-Au300-R12/13, Zhenjiang Lehua Technology Co. Ltd., China) and blotted for 3 s under a blot force of 0 at 100% humidity and 16C before being flash-frozen in liquid ethane with a Mark IV Vitrobot system (FEI). Micrographs were acquired on a Titan Krios microscope (FEI) operated at 300 kV with a K2 Summit direct electron detector (Gatan). SerialEM was used for automatic data collection. A nominal magnification of 22,500 was used for imaging, which yielded a pixel size of 1.04 . The defocus range was between 1.5 and 2.5 m. Each micrograph was dose-fractionated to 32 frames under a dose rate of 9.4 e/2 per second and an exposure time of 6.4 s, which resulted in a total dose of about 60 e/2.

Motion correction and exposure weighting was performed by the MotionCorr2 program (35), and the CTF (contrast transfer function) parameter was estimated using the Gctf program (36). The automatic particle picking was performed by Gautomatch (developed by K. Zhang, MRC Laboratory of Molecular Biology, Cambridge, UK) and Auto-picking module in RELION; an initial model was made by e2initialmodel.py in EMAN2 software package (37), and all other steps were performed using RELION (38). For the air-oxidized ACIII dataset, 600 particles were manually picked and extracted for two-dimensional (2D) classification. The resulting 2D class averages were used as the templates for the automated particle picking, which yielded 257,815 particles from 1700 micrographs. The picked particles were extracted at 2 2 binning and subjected to three rounds of 2D classification. A total of 197,496 particles were finally selected for 3D classification.

Good 2D class averages in different orientations were selected to generate the initial model. A total of 177,489 particles were left after two rounds of 3D classification and re-extracted into the original pixel size of 1.04 . The following 3D refinement and postprocessing yielded an EM map with a resolution of 3.45 . After performing CTF refinement in RELION3, the resolution was increased to 3.24 . Reported resolutions were estimated with a soft-edge mask around the protein complex and micelle densities and based on the gold-standard FSC (Fourier Shell Correlation) = 0.143 criterion. Local resolution was estimated with Resmap (39).

For the reduced ACIII dataset, 1970 unscreened micrographs were subjected to 3D referencebased auto-picking in RELION3; reconstruction of the ACIII dataset was the 3D reference low passfiltered to 20 . The resulting 488,581 particles were used to extract particles at 2 2 binning. After two rounds of 2D classification, 297,122 particles were selected for a 3D refinement and alignment-free 3D classification, and 219,913 particles from the best 3D class were re-extracted without downscaling. The following 3D refinement and postprocessing yielded an EM map with a resolution of 3.68 . CTF refinement and another alignment-free 3D classification improved the resolution to 3.51 and 3.46 , respectively. The final subset had 207,633 particles.

De novo atomic model building was conducted in Coot (40). Sequence assignments were guided by residues with bulky side chains. The starting models of the cofactors were taken from the CCP4 ligand library. The model was real spacerefined by PHENIX (41, 42) with intra-cofactor and protein-cofactor geometric constraints. The refinement and model statistics are listed in Table 1. All figures were prepared in PyMOL (www.pymol.org) or UCSF Chimera (43).

The difference map between air-oxidized and dithionite-reduced ACIII was calculated using EMAN2 (37). First, the cryo-EM map of dithionite-reduced ACIII was fitted to that of air-oxidized ACIII by Chimera and then was clipped into the same box size using e2proc3d.py in EMAN2. Then, the structural amplitudes of both maps were scaled using e2proc3d.py in EMAN2. Last, the difference map between the corrected maps was computed by the e2.py python tool in EMAN2 and further low-passfiltered at a quarter of the Nyquist criterion.

The rest is here:
Cryo-EM structures of the air-oxidized and dithionite-reduced photosynthetic alternative complex III from Roseiflexus castenholzii - Science Advances

NEW YORK, July 28, 2020 /PRNewswire/ -- According to the current analysis of Reports and Data, theProtein Engineering marketwas valued at USD 1.86 billion in 2019 and is expected to reach USD 4.77 billion by the year 2027, at a CAGR of 12.3%. Protein engineering is the process of conception and production of unnatural polypeptides, which is achieved through the modification of different amino acid sequences that are found in nature. With the wide application of protein engineering, various synthetic protein structures and functions can now be designed completely using a computer and produced in the laboratory using various methods.

With the advent of technology, the rising demand is expected to provide traction to the market. For instance, computational protein design holds a potential promise to revolutionize protein engineering. Protein engineering is significantly in demand as it can develop useful or valuable proteins. It is known to be an emerging research field that helps researchers in understanding protein folding and recognition used for protein design principles. Protein engineering market is spurred by various reasons such as availability of immense information regarding 3D protein structure, advancement in structural bioinformatics, novel protein design algorithms, and other factors that have made it possible to use computational approaches for research and development in protein engineering. Strategic utilization of approaches that have been discovered with research has enabled better stability and increased catalytic activity of the protein. Moreover, it has increased its applicability in various fields that will further expand the market. Protein engineering has also evolved to become a potent tool contributing considerably to the developments in both synthetic biology and metabolic engineering. The rising funding for synthetic biology by governments and other healthcare institutions may drive the industry extensively in the future. Furthermore, substantial ongoing research in the drug discovery process is expected to fuel the market during the forecast period. Growing technological advancements and innovative instruments useful for protein engineering may drive the market extensively in the future. The emergence of new diseases due to microorganisms may also trigger market growth.

Request free sample of this research report at: https://www.reportsanddata.com/sample-enquiry-form/3369

However, factors such as high cost of instrumentation and lack of skill professional may hamper the protein engineering industry in the forecast period.

COVID-19 Impact:

WHO is focused on the latest scientific findings and knowledge on COVID-19. Various researchers and scientists are discovering new ways to tackle COVID-19 infection. Furthermore, the market of protein engineering is also going to get significantly affected due to the multiple kinds of research which are being carried out in this field. With the advent of technology, improving protein stability is an essential goal for clinical and industrial applications, however no commonly accepted and widely used strategy for efficient engineering is known. Furthermore, during the COVID-19 outbreak, protein engineering market is significantly impacted because many of the procedures involve lab automation. Market players are focusing on a research-based approach that will provide traction to the market. For instance, Vir Biotech identified two antibodies that could be effective in preventing and treating COVID-19. However, the high requirement of funds is expected to hamper the market during this period.

To identify the key trends in the industry, click on the link below:https://www.reportsanddata.com/report-detail/protein-engineering-market

Further key findings from the report suggest

Order Now: https://www.reportsanddata.com/checkout-form/3369

For the purpose of this report, Reports and Data has segmented the Protein Engineering market on the basis of product, type, technology, end use and region:

By Product Outlook (Revenue in Million USD; 2017-2027)

By Type Outlook (Revenue in Million USD; 2017-2027)

By Technology Outlook (Revenue in Million USD; 20172027)

By End Use Outlook (Revenue in Million USD;20172027)

Regional Outlook (Revenue, USD Billion; 2017-2027)

Browse more similar reports on Biotechnology category by Reports And Data

About Reports and Data

Reports and Data is a market research and consulting company that provides syndicated research reports, customized research reports, and consulting services. Our solutions purely focus on your purpose to locate, target and analyze consumer behavior shifts across demographics, across industries and help client's make a smarter business decision. We offer market intelligence studies ensuring relevant and fact-based research across a multiple industries including Healthcare, Technology, Chemicals, Power and Energy. We consistently update our research offerings to ensure our clients are aware about the latest trends existent in the market. Reports and Data has a strong base of experienced analysts from varied areas of expertise.

Contact Us:

John WHead of Business DevelopmentReports And Data | Web:www.reportsanddata.comDirect Line: +1-212-710-1370E-mail: [emailprotected]LinkedIn | Twitter | Blogs

SOURCE Reports And Data

The rest is here:
Protein Engineering Market To Reach USD 4.77 Billion By 2027 | Reports And Data - PRNewswire

- The increasing research in protein engineering and rising government funding for new drug discovery are expected to be some major drivers for the market

- Market Size USD 1.86 Billion in 2019, Market Growth CAGR of 12.3%, Market Trends The growing research institutions across developing economies is expected to fuel the market in the forecast period

NEW YORK, July 28, 2020 /PRNewswire/ -- According to the current analysis of Reports and Data, theProtein Engineering marketwas valued at USD 1.86 billion in 2019 and is expected to reach USD 4.77 billion by the year 2027, at a CAGR of 12.3%. Protein engineering is the process of conception and production of unnatural polypeptides, which is achieved through the modification of different amino acid sequences that are found in nature. With the wide application of protein engineering, various synthetic protein structures and functions can now be designed completely using a computer and produced in the laboratory using various methods.

With the advent of technology, the rising demand is expected to provide traction to the market. For instance, computational protein design holds a potential promise to revolutionize protein engineering. Protein engineering is significantly in demand as it can develop useful or valuable proteins. It is known to be an emerging research field that helps researchers in understanding protein folding and recognition used for protein design principles. Protein engineering market is spurred by various reasons such as availability of immense information regarding 3D protein structure, advancement in structural bioinformatics, novel protein design algorithms, and other factors that have made it possible to use computational approaches for research and development in protein engineering. Strategic utilization of approaches that have been discovered with research has enabled better stability and increased catalytic activity of the protein. Moreover, it has increased its applicability in various fields that will further expand the market. Protein engineering has also evolved to become a potent tool contributing considerably to the developments in both synthetic biology and metabolic engineering. The rising funding for synthetic biology by governments and other healthcare institutions may drive the industry extensively in the future. Furthermore, substantial ongoing research in the drug discovery process is expected to fuel the market during the forecast period. Growing technological advancements and innovative instruments useful for protein engineering may drive the market extensively in the future. The emergence of new diseases due to microorganisms may also trigger market growth.

Request free sample of this research report at: https://www.reportsanddata.com/sample-enquiry-form/3369

However, factors such as high cost of instrumentation and lack of skill professional may hamper the protein engineering industry in the forecast period.

COVID-19 Impact:

WHO is focused on the latest scientific findings and knowledge on COVID-19. Various researchers and scientists are discovering new ways to tackle COVID-19 infection. Furthermore, the market of protein engineering is also going to get significantly affected due to the multiple kinds of research which are being carried out in this field. With the advent of technology, improving protein stability is an essential goal for clinical and industrial applications, however no commonly accepted and widely used strategy for efficient engineering is known. Furthermore, during the COVID-19 outbreak, protein engineering market is significantly impacted because many of the procedures involve lab automation. Market players are focusing on a research-based approach that will provide traction to the market. For instance, Vir Biotech identified two antibodies that could be effective in preventing and treating COVID-19. However, the high requirement of funds is expected to hamper the market during this period.

To identify the key trends in the industry, click on the link below:https://www.reportsanddata.com/report-detail/protein-engineering-market

Further key findings from the report suggest

Order Now: https://www.reportsanddata.com/checkout-form/3369

For the purpose of this report, Reports and Data has segmented the Protein Engineering market on the basis of product, type, technology, end use and region:

By Product Outlook (Revenue in Million USD; 2017-2027)

By Type Outlook (Revenue in Million USD; 2017-2027)

By Technology Outlook (Revenue in Million USD; 20172027)

By End Use Outlook (Revenue in Million USD;20172027)

Regional Outlook (Revenue, USD Billion; 2017-2027)

Browse more similar reports on Biotechnology category by Reports And Data

About Reports and Data

Reports and Data is a market research and consulting company that provides syndicated research reports, customized research reports, and consulting services. Our solutions purely focus on your purpose to locate, target and analyze consumer behavior shifts across demographics, across industries and help client's make a smarter business decision. We offer market intelligence studies ensuring relevant and fact-based research across a multiple industries including Healthcare, Technology, Chemicals, Power and Energy. We consistently update our research offerings to ensure our clients are aware about the latest trends existent in the market. Reports and Data has a strong base of experienced analysts from varied areas of expertise.

Contact Us:

John WHead of Business DevelopmentReports And Data | Web:www.reportsanddata.comDirect Line: +1-212-710-1370E-mail: sales@reportsanddata.comLinkedIn | Twitter | Blogs

Logo: https://mma.prnewswire.com/media/1009077/Reports_And_Data_Logo.jpg

SOURCE Reports And Data

View post:
Protein Engineering Market To Reach USD 4.77 Billion By 2027 | Reports And Data - PR Newswire UK