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Why does the inhibition of translation initiation cause the accumulation of 80S ribosomal monosomes?

Why does the inhibition of translation initiation cause the accumulation of 80S ribosomal monosomes?



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As I have read in (1), inhibition of translation initiation will increase the number of 80S ribosomes while decreasing the fraction of polysomes due to the polysome-runoff. The net effect is to decrease the polysome/monosome (P/M) ratio and we observe it for example by inducing ER stress by tunicamycin in (2).

I understand the reduction in polysomes - since we don't interefere with elongation, polysomes work as normal, fallling of eventually, but new monosomes cannot proceed to polysomal stage due to inhibited initation.

However, why do the 80S ribosomes accumulate? I have always thought that 80S ribosome is always bound to mRNA, otherwise it dissociates as separate 40S and 60S subunits. Does this effect therefore mean that the accumulated 80S fraction is mRNA-free and therefore "vacant 80S ribosomes", as decribed in (1), can exist?

Thanks a lot!

  1. https://books.google.co.uk/books?id=6sEq1xLu_hcC&pg=PA130&lpg=PA130&dq=polysome+translation+initiation&source=bl&ots=Un6JjImBJX&sig=ln9mn0KoJDJaVOGK8LbrEmMz9XE&hl=cs&sa=X&ei=x2JiVci8N4jX7Ab02YLgBQ&ved=0CGQQ6AEwCw#v=onepage&q=polysome%20translation%20initiation&f=false

  2. http://jcs.biologists.org/content/115/11/2443/F4.large.jpg">


    It will depend on the drug you are using. There is a pathway that forms the translational initiation complex that starts with binding the mRNA's cap, the small subunit scans to find the AUG, then the large subunit binds, and so on. The 80 S peaks on the density gradients are not vacant, they are poised on mRNAs waiting to initiate (if you removed the inhibitor).


    Mechanism of action of streptomycin in E. coli: interruption of the ribosome cycle at the initiation of protein synthesis

    MECHANISM OF ACTION OF STREPTOMYOCIN IN E. COLI: INTERAPTION OF THE RIBOSOME CYCLE AT THE INITATION OF PROTEIN SYNTHESIS.

    There are several kind of antibiotics with a different antimicrobial mechanism which can be used against gram positive and negative bacteria. There are also many kinds of classifications can be used in order to classify the antibiotics. Some of these antibiotics have a broad-spectrum bacterial effect whereas others have narrow- spectrum bacterial effect. Some of them have bactericidal (they kill the bacteria) impact and others have bacteriostatic (they inhibit the bacterial growth or replication) impact. Some antibiotics can be both bactericidal and bacteriostatical.

    Different antibiotics have different modes of action and target sites within bacterial cells. There are five basic mechanisms of antibiotic action against bacterial cells

    • Inhibition of cell wall synthesis
    • Inhibition of protein synthesis (translation)
    • Alteration of cell membranes
    • Inhibition of nucleic acid synthesis
    • Antimetabolite activity

    Inhibition of cell wall synthesis is the most common mechanism of antibiotics. Second largest class antibiotics are showing their antimicrobial effect by inhibiting the translation mechanism in the cell. In the paper of Luzzatto and colleagues, they proved the antimicrobial mechanism of streptomycin on Escherichia coli by the paper published on 1968.

    There are a couple of ways of antibiotics inhibiting translation in the cell. These are

    1. tRNA mimicry
    2. Inhibitors of peptide-bone formation
    3. Inhibitors of binding of tRNA to the A site
    4. Inhibitors of translocation
    5. Binding to 23S RNA
    6. Binding to the 30S ribosome

    Streptomycin belongs to aminoglycoside class antibiotic and it is known as a protein synthesis inhibitor.

    Back at 1964, in a study called “Streptomycin, Suppression and the Code” conducted by Julian Davies and Walter Gilbert, they were aware of the protein synthesis inhibiting the power of streptomycin, however, the mechanism was unknown. Previous studies of this study had suggested that streptomycin blocked the protein synthesis by strongly binding to nucleic acids. Julian Davies had concluded that streptomycin had done some alterations in the coding properties. Moreover provided the evidence that the ribosomes control the accuracy of the reading and may have a role in suppression. After that study, in 1968 Luzzatto lightened the unknown mechanism of streptomycin inhibiting the translation by this research.

    In the paper of Luzzatto, they found that a certain concentration (lethal concentration) of streptomycin cause the accumulation of 70S ribosomes in E. coli cells. These ribosomes are incapable of doing translation in the cell. These 70S ribosomal units that accumulated were called “streptomycin monosomes”. Moreover, they found that these units consist of a complex of 30S and 50S subunits, tRNA, mRNA, and streptomycin. They observed that the 70S subunits which consist streptomycin had abnormal initiation complexes that cannot elongate therefore they accumulate. So, they conclude that streptomycin molecules somehow “freeze” the protein initiation in E. coli.

    According to their findings Streptomycin blocks bacterial protein synthesis at initiation. After intact bacteria are exposed to streptomycin, polysomes become rapidly depleted and 70S particles. “The streptomycin monomers” build up. Although the formation of initiation complex is not affected, the complex formed in the presence of streptomycin cannot synthesize protein and remains fixed in the position. It is proposed that ribosome beyond the initiation stage are able to continue their movement and detachment so that a 70S ribosomal complex of mRNA and 50S and 30S units with bound streptomycin results. In effect, the initiation complex is frozen.

    In their experiment, they used Escherichia coli mutant sud 24 and grow it in fragile form to be able to lyse and analyze during the translations. To trace the RNA during translations they used radioactively labelled RNAs. They measured the speed of the molecules by using sucrose gradient analysis to determine the size distribution.

    They used a streptomycin resistant derivative (N21) and susceptible AB301 strain, labelled ala-transfer tRNA, and labeled F-met tRNA, natural mRNA (f-met dependent), synthetic mRNA (poly AUG, f-met independent) and also they used a phage protein called R17.

    At the moment they add streptomycin the ribosome was the 30S and the 50S form as well as bound to mRNA. After 20 and 40 min intervals they released (a) a decrease in large polyribosomes and in free 30S and 50S particles. (b) Accumulation of 70S monomers. (fig.1)

    The protein synthesis directed by the phage R17 was observed alone with streptomycin and streptomycin was observed alone without the phage R17. Their finding had shown that streptomycin blocked the function of the natural mRNA (fig. 2)

    Streptomycin blocked the RNA synthesis directed by natural mRNA but not with the synthetic mRNA. The natural mRNA initiates the protein synthesis by using f-met however the synthetic mRNA doesn’t require the f-met to start the translation. Thereby, they released that the streptomycin was blocking somehow the f-met boundary on the 30S ribosomal subunit to and block the F-met to come and attached in order to initiate the translation process. However, the synthetic RNA experiment did not block by adding the streptomycin. The amount of S 35 f-met-tRNA was reduced by streptomycin and H 3 -ala-tRNA bound to ribosomes.

    They realized that the initiated protein synthesis with 70S ribosomes was not blocked and continue when streptomycin added but only new translation processes did not initiate. So, by using the synthetic mRNA and natural and also by using a radioactive label with the knowledge of f-met requirement for the translation they were able to understand this mode of action of streptomycin was at the initiation right after ribosomal subunits association.


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    MATERIALS AND METHODS

    Affinity cross-linking of s 4 U-containing mRNA analogues to human ribosomes

    Human ribosomal 40S and 60S subunits were isolated from full term placenta, as described in ( 23). Purified tRNA Phe (∼80%) and tRNA Val (∼70%) from Escherichia coli were the kind gifts from Dr V.I. Katunin (St. Petersburg Nuclear Physics Institute named by B.P. Konstantinov of National Research Center ‘Kurchatov Institute’, Gatchina, Russia). Yeast tRNA Asp transcript was obtained by T7 transcription in vitro, as described ( 24). DNA templates for the synthesis of mRNA analogues with randomly distributed s 4 U residues by T7 transcription in vitro were obtained by hybridization of the following oligodeoxyribonucleotide pairs: F-Asp, 5′-aaattaatacgactcactatagggaagaaagaagataaagaaaaagaa-3′ and R-Asp, 5′-ttctttttctttatcttctttcttccctatagtgagtcgtattaattt-3′ F-PheAsp, 5′-aaattaatacgactcactatagggaagaaagaattcgataaagaaaaa-3′ and R-PheAsp, 5′-tttttctttatcgaattctttcttccctatagtgagtcgtattaattt-3′ F-PheVal, 5′-aaattaatacgactcactatagggaagaaagaattcgtaaaagaaaaa-3′ and R-PheVal, 5′-tttttcttttacgaattctttcttccctatagtgagtcgtattaattt-3′ F-PheVal(C-rich), 5′-cgattaatacgactcactatagggaagccaccattcgtacaccaccac-3′ and R-PheVal(C-rich), 5′-gtggtggtgtacgaatggtggcttccctatagtgagtcgtattaatcg-3′. The T7 transcription reaction was carried out as described ( 25) the concentrations UTP and s 4 UTP in reaction mixture were 0.5 mM. After the reaction, the synthesized RNAs were purified by 12% denaturing PAGE and used as mRNA analogues. There were 5′-GGGAAGAAAGAAGAs 4 UAAAGAAAAAGAA-3′, 5′-GGGAAGAAAGAAs 4 Us 4 UCGs 4 UAAAAGAAAAA-3′, 5′-GGGAAGAAAGAAs 4 Us 4 UCGAs 4 UAAAGAAAAA-3′ and 5′-GGGAAGCCACCAs 4 Us 4 UCGs 4 UACACCACCAC-3′ designated hereinafter as mRNA I, II, III and IV, respectively. If necessary, the mRNAs and tRNAs were 5′ end dephosphorylated with FastAP alkaline phosphatase (Thermo Scientific) and then 5′ end 32 P-labeled in reaction with [γ- 32 P]ATP and T4 polynucleotide kinase. Complexes of 80S ribosomes with mRNAs and tRNAs with codon-anticodon interactions either at the P site or at the P and A sites simultaneously, were obtained according to ( 12). The levels of binding to 80S ribosomes of 32 P-labeled tRNA Asp or tRNA Phe cognate to an mRNA triplet targeted to the P-site to form a ternary complex or to the A site to convert the latter into a quaternary one, and of the respective 32 P-labeled mRNAs were measured by nitrocellulose filtration assay as described ( 12). For cross-linking in the ternary complexes with mRNAs I-IV, reaction mixtures included 80S ribosomes (0.83 μM), 32 P-labeled mRNA (2 μM), and the tRNA (7 μM) in 50 mM Tris–HCl (pH 7.5) containing 100 mM KCl, 13 mM MgCl2 and 0.5 mM EDTA (buffer A). For cross-linking in the quaternary complexes with mRNAs II and III, the respective reactive mixtures were supplemented with the appropriate tRNA to its concentration of 13 μM. After incubation under binding conditions ( 12), the above mixtures were irradiated with mild UV light (λ > 300 nm) ( 26), followed by analysis of cross-linked ribosomal proteins in 12% SDS PAGE as described ( 12). The identification of the cross-linked ribosomal proteins was carried out based on the retardation effect of the cross-linked RNA fragments on the electrophoretic mobility of ribosomal proteins known from our previous works (e.g. see ( 7–9, 12, 14)).

    Cells culturing and transfection

    The minigene of FLAG uS19 amplified with the use of HEK293 cDNA and primers F (5′-acgtgaattcatggtggactacaaagacgatgacgacaaggcagaagtagagcagaag-3′) and R (5′- acgtgaattcttacttgagagggatgaagc-3′) was inserted into pAG-1 vector ( 26) at EcoRI site in the forward direction. The resulting plasmid pAG-1( FLAG uS19) was linearized by BamHI and then utilized to transfect HEK293 cells (ATCC CRL-1573) using Lipofectamin LTX (Invitrogen). The selection of cell clones stably producing the target protein was performed as described ( 26). Stable clones were tested for their ability to produce FLAG uS19 after doxycycline (DOX) induction by western blotting using anti-FLAG M5 (Sigma, #F4042) as described ( 26).

    PAR-CLIP analysis of a stable cell line producing FLAG uS19

    The PAR-CLIP procedure was carried out according to ( 22) with some modifications in three biological replicates. Typically, adherent pAG-1( FLAG uS19)-derivatized HEK293 cells in four 15-cm Petri dishes were cultured in Dulbecco's modified Eagle's medium (DMEM) containing 10% FBS and 100 U/ml of Penicillin–Streptomycin (all from Thermo Fisher Scientific) in CO2-incubator (5% CO2) at 37°C. At 60–70% confluency, the production of FLAG uS19 in cells was induced with DOX (2 μg/ml) and after 44 h, s 4 U was added to 250 μM. After 6 h, cycloheximide was added to a concentration of 100 μg/ml and the cells were kept on ice for 10 min, followed by washing with ice-cold PBS. Cross-linking was performed by irradiation of the cells with UV light at 365 nm (0.5 J/cm 2 ) in Bio-Link (Vilber Lourmat) on ice. In the control experiment, s 4 U was not added to the cell medium and the cells were not UV-irradiated. Finally, the cells were harvested with ice-cold PBS and centrifuged at 1000 g.

    The cell pellet was lysed in three volumes of lysis buffer (20 mM Tris–HCl (pH 7.5), 150 mM NaCl, 5 mM MgCl2, 1mM DTT, 1% Triton-X100, 100 μg/ml of cycloheximide and 0.02 U/μl of Turbo DNase I (Ambion)) for 10 min on ice and then triturated through 29 G needle 10 times. The cell lysate was clarified by centrifugation at 20 000 g for 15 min at 4°C and the supernatant was transferred into a new Eppendorf tube. RNase I (Ambion) was added to 1.3 U/μl, and the reaction mixture was incubated at ambient temperature for 45 min, followed by the addition of SUPERase In RNase inhibitor (Thermo Fisher Scientific) to 0.35 U/μl. The mixture was then clarified as described above, and the supernatant (∼750 μl) was layered onto 1 ml of 30% sucrose cushion (containing 20 mM Tris–HCl (pH 7.5), 150 mM NaCl, 5 mM MgCl2, 1 mM DTT, 1% Triton-X100, 100 μg/ml of cycloheximide and 0.02 U/μl of SUPERase In) in a SW60 rotor centrifuge tube. Free tube volume was supplemented with 20 mM Tris–HCl (pH 7.5) buffer containing 150 mM NaCl, 5 mM MgCl2 and 1 mM DTT, and the tube was centrifuged in SW60 Ti rotor at 25 000 rpm for 16 h at 4°C. The pellet was dissolved in 10 mM ethylenediaminetetraacetic acid (EDTA) containing 1% SDS, incubated for 30 min at 37°C and then diluted 20-fold with IP buffer (20 mM Tris–HCl (pH 7.5), 150 mM NaCl, 0.1% Nonidet P40 and protease inhibitor cocktail (Sigma)). The mixture was clarified as described above, and anti-FLAG M2 antibodies (Sigma, #F1804) bound to magnetic Protein G beads (Dynabeads, Life Technologies) were added to the supernatant. Typically, 10 μg of antibodies were bound to 40 μl of Dynabeads in one experiment as described ( 26). Immunoprecipitation was carried out for 3 h at 4°C on a rotating platform and the supernatant was then removed. The beads were washed with 1 ml of IP buffer (50 mM Tris–HCl (pH 7.5), 300 mM NaCl, 0.05% Nonidet P40 and 0.5 mM DTT), three times with 1 ml of HS buffer (50 mM Tris–HCl (pH 7.5), 500 mM NaCl, 0.05% Nonidet P40 and 0.5 mM DTT) and twice with 1ml of Defos buffer (50 mM Tris–HCl (pH 7.5), 100 mM NaCl, 10 mM MgCl2 and 1 mM DTT). The remaining procedures, including the dephosphorilation of RNA fragments cross-linked to FLAG uS19, their 5′-end labeling with [γ- 32 P]ATP and phosphorylation with ATP as well as cross-link separation by SDS-PAGE followed by transfer onto the nitrocellulose membrane and the isolation of cross-linked RNA fragments, were carried out as described ( 26).

    NGS sequencing, data processing and bioinformatics analysis

    DNA libraries were prepared according to ( 27) and sequenced with 2 × 75 bp paired-ends reagents on MiSeq (Illumina) in SB RAS Genomics Core Facility (Institute of Chemical Biology and Fundamental Medicine, SB RAS, Novosibirsk, Russia). Reads were demultiplexing by CEL-Seq script ( 28). Adapter and low-quality sequences were removed using TrimGalore v.0.5.0 software (http://www.bioinformatics.babraham.ac.uk/projects/trim_galore). Read mapping was performed against the Ensembl annotated human genome (GRCh38.93) with STAR v.2.6.1 software ( 29). BAM deduplication by unique molecular indexes was carried out with ‘dedup’ script from UMI-tools ( 30). Transitions were found by Basic Variant Detection in CLC GW v.12.0 software (Qiagen) and filtered by custom python scripts. The read data reported in this study were submitted to the GEO under the accession number GSE128265.

    The uS19 cross-linking sites were detected by the appearance of characteristic T/C (for plus DNA stands) and A/G (for minus DNA strands) transitions in the sequencing reads, which were found as described above, with parameters: minimum coverage, 5 minimum count, 1 minimum frequency, 1%. The resulting variant call format (VCF) files were used for subsequent downstream analysis that was carried out using R Bioconductor platform packages. Genomic positions from VCF files were used as coordinates of cross-linking sites and were annotated by ensembl transcript IDs and HGNC gene symbols using ‘biomaRt’ package ( 31) and Ensembl annotation GRCh38.93. Coordinates of transitions that corresponded to the non-protein coding part of the genome and the mitochondrial genome, were removed from the main part of the subsequent analysis. Genomic positions were transformed to internal CDS coordinates according to Ensembl annotation GRCh38.93. The resulting peaks were identified using the ‘chipseq’ and ‘coverage view’ packages and were matched with the positions of the above-mentioned transitions. Only peaks overlapping with transition positions and having coverage above the threshold were considered as read clusters. Among the transition positions that corresponded to a particular cluster, only the position with the highest T/C transition coverage was considered as the main cross-linking site, where appropriate.

    CDS sequences were downloaded from Ensembl (www.ensembl.org/info/data/ftp/Homo_sapiens.GRCh38.cds.all.fa) and subsequences from –19 triplet upstream to +19 triplet downstream of the transition internal CDS coordinate were extracted. Considering the open reading frames, the resulting sequences were translated into amino acid sequences that were used for the LOGO analysis utilizing the Weblogo tool (https://weblogo.berkeley.edu/). The read clusters were sorted based on the total cluster read coverage, and the transition context LOGOs were created for all found clusters, including the 100 most covered ones (‘top100’). Histograms and graphics were built using integrated R tools.

    The correlation between PAR-CLIP replicates was measured using the standard R function (cor), and the resulting plots were generated using the ggplot2 package. To compare the raw reads to each other, read count correlation tables were generated using the summarizeOverlaps function with the default parameters from the GenomicAlignment package ( 32). For all identified clusters, the total coverage was estimated using the CoverageView package, and then applied to correlation measurements.

    To find proteins with (E/K)12 motifs, a set of all human protein sequences was recovered from Ensembl (https://www.ensembl.org/info/data/ftp/). The amino acid sequences were scanned for the presence of any of the 2 12 (E/K)12 motifs using the Bioconductor Biostrings package with the vcount-Pattern function with the max.mismatch parameter equal to 4 and other parameters default.

    Ribosome profiling with cells producing FLAG uS19 and cells grown in a medium containing s 4 U

    For ribosome profiling with HEK293 cells producing FLAG uS19, pAG-1( FLAG uS19)-derivatized HEK293 cells were grown in three biological replicates as in the PAR-CLIP experiment. For the same task with HEK293 cells grown in the presence of s 4 U, the cells were cultured in 10-cm Petri dishes as described above in three biological replicates up to 60% confluency, and then s 4 U was added to one half of the Petri dishes up to 250 μM, followed by incubation for 16 h and the cells collection. Subsequent manipulations, including cell lysis, RNase I treatment of the lysate and its centrifugation through a sucrose cushion were the same as described above. With HEK293 cells producing FLAG uS19, the resulting pellet was solubilized in 200 μl of buffer A and incubated with 15 μl of Protein G beads pre-bound with anti-FLAG M2 antibodies for 3 h at 4°C followed by separation of the supernatant from the beads. Total RNA was isolated from the supernatant fraction containing ribosomes with endogenous uS19 and from ribosomes remaining on beads and containing uS19 FLAG using TRIzol (Ambion) according to the manufacturer's protocol. Total RNA from ribosomal pellet obtaining after centrifugation of the cell lysate through a sucrose cushion in the experiment with HEK293 cells grown in the presence of s 4 U was isolated in the same way. Subsequent procedures, including RNA fragments size selection and purification, were performed according to ( 33). DNA libraries were prepared and sequenced as above. The corresponding Ribo-seq data were submitted to the GenBank under the study accession PRJNA563539 and the sample accession SRP220276.


    Abstract

    Gene expression is highly accurate and rarely generates defective proteins. Several mechanisms ensure this fidelity, including specialized surveillance pathways that rid the cell of mRNAs that are incompletely processed or that lack complete open reading frames. One such mechanism, nonsense-mediated mRNA decay, is triggered when ribosomes encounter a premature translation-termination — or nonsense — codon. New evidence indicates that the specialized factors that are recruited for this process not only promote rapid mRNA degradation, but are also required to resolvea poorly dissociable termination complex.


    MATERIALS AND METHODS

    Overview of the protocol

    Polysome profiling separates translated mRNAs on a sucrose gradient according to the number of bound ribosomes. First, cells are lysed and loaded on top of a 15–40% sucrose gradient. After ultracentrifugation, the gradient is monitored at A254 using a flow cell coupled to a spectrophotometer and then fractionated into equal fractions: untranslated mRNAs (top fractions) are separated from polysome-associated mRNAs (bottom fractions). Fractions are then processed for RNA extraction, either manually by acid phenol–chloroform extraction or in a 96-well format using an automated RNA purification processor, simultaneously handling several gradients. The translational status of a given mRNA species is analyzed by RT-PCR amplification and its relative quantification in each fraction. Alternatively, the content of the polysomal fractions can be identified using a global analysis (such as high-throughput sequencing), granting access to the cell translatome. A schematic overview of the method is presented in Figure 1.

    Overview of the polysome profiling protocol to analyze translation activity. The various steps of the protocol involve (1) cell lysis, (2) sucrose-gradient centrifugation and (3) fractionation, (4) RNA extraction and RNA integrity check, (5) analysis of translational status of mRNAs. See the text for details.

    Overview of the polysome profiling protocol to analyze translation activity. The various steps of the protocol involve (1) cell lysis, (2) sucrose-gradient centrifugation and (3) fractionation, (4) RNA extraction and RNA integrity check, (5) analysis of translational status of mRNAs. See the text for details.

    Detailed protocol

    Sample collection

    Experiments were performed using Paracentrotus lividus or Sphaerechinus granularis sea urchins the experiments shown herein correspond to P. lividus samples. Sea urchins were collected in the bays of Crozon or Concarneau (Brittany, France). Gametes were obtained by intracoelomic injection of 1 ml 0.1 M acetylcholine. Eggs were collected in filtered seawater (FSW), filtered on hydrophilic gauze and washed twice in FSW by centrifugation for 2 min at 2000 rpm (Heraeus, Labofuge 400 with swinging buckets). Eggs were incubated 40 s in FSW supplemented with 0.7 mM citric acid to remove the jelly coat, and rinsed again in FSW. For fertilization, eggs were suspended in FSW as a 5% suspension. Sperm was collected in a Petri dish and stored at 4°C until use. Extemporaneous dilution in FSW (10 μl of sperm in 1 ml of FSW) activates sperm for fertilization and 10 μl of this diluted sperm was added per ml of egg suspension. Embryos were cultured at 16°C under constant agitation. Only batches of embryos displaying a fertilization and division rate above 95% were used.

    Cell lysis

    Eggs or embryos were collected in FSW by brief centrifugation (1 min at 1000 rpm, Heraeus, Labofuge 400 with swinging buckets), and the pellet was resuspended in four volumes of cold lysis buffer (10 mM Tris pH 7.4 250 mM KCl 10 mM MgCl2 25 mM ethylene glycol-bis(2-aminoethylether)-N,N,N',N'- tetraacetic acid (EGTA) 0.4% Igepal 5% sucrose RNase-free water and extemporaneously 1 mM 1,4-Dithiothreitol (DTT) 10 μg/ml aprotinin 2 μg/ml leupeptin 100 μM emetine 40 U/ml RNase inhibitor). Lysis was done in a Dounce homogenizer using 10 strokes of the tight B pestle. All steps were done at 4°C, on ice or in the cold room. The lysate was then centrifuged for 10 min at 13000 rpm in a tabletop centrifuge to remove nuclei and cellular debris. The supernatant was carefully transferred into a new microtube. Samples can be frozen in liquid nitrogen and kept at −80°C until further use for polysome fractionation.

    The concentration of nucleic acid in the lysate was measured by absorbance at A260nm of a 5 μl sample of lysate diluted in 1 ml of water using a spectrophotometer. Starting from 10 6 eggs or embryos in a 250 μl pellet, typical yield for sea urchin samples was usually between 20 and 40 ODA260.

    Critical steps for lysate preparation

    The protocol described above was defined after optimizing several parameters such as starting material (frozen or fresh eggs), lysis buffer composition and volume and lysis techniques. Optimization of the protocol for sea urchin oocytes and embryos is detailed below, and the quality of the RNA in the lysate was tested for each variant of the protocol using acid phenol–chloroform extraction and electrophoresis. We also provide suggestions for adapting this protocol to other models and organisms.

    We observed that RNA quality improved when the lysates were prepared with fresh eggs rather than from a frozen egg pellet kept at −80°C (Figure 2, lanes A and B). Alternatively, freezing and grinding under liquid nitrogen prior to lysis works for organisms with cell walls or for solid tissues ( 12, 25). Lysates can be kept frozen at −80°C until use, without detectable loss of RNA integrity.

    RNA quality with different conditions for lysis of sea urchin eggs. Lysis was done using a 25G needle (A–E) or a Dounce homogenizer (F–J), on frozen eggs (A) or fresh eggs (B–J). The same volume (V) of eggs was lysed in increasing volumes of lysis buffer ranging from 2:1 to 1:4 (B–E). An additional wash with filtered seawater FSW (G) or Ca 2+ -free SW (H) was tested before lysis. Lysis buffer contained either 1 mM EDTA (F) or 25 mM EGTA (I and J). Two RNA quantities prepared using the optimized protocol showed only the 28S and 18S RNA without degradation products (lane I: 250 ng lane J: 1 μg). RNAs from each lysate were obtained after an acid phenol–chloroform extraction, and separated on a 2% agarose-TBE gel to check for integrity.

    RNA quality with different conditions for lysis of sea urchin eggs. Lysis was done using a 25G needle (A–E) or a Dounce homogenizer (F–J), on frozen eggs (A) or fresh eggs (B–J). The same volume (V) of eggs was lysed in increasing volumes of lysis buffer ranging from 2:1 to 1:4 (B–E). An additional wash with filtered seawater FSW (G) or Ca 2+ -free SW (H) was tested before lysis. Lysis buffer contained either 1 mM EDTA (F) or 25 mM EGTA (I and J). Two RNA quantities prepared using the optimized protocol showed only the 28S and 18S RNA without degradation products (lane I: 250 ng lane J: 1 μg). RNAs from each lysate were obtained after an acid phenol–chloroform extraction, and separated on a 2% agarose-TBE gel to check for integrity.

    The ratio of the volume of the egg pellet to lysis buffer was optimized in our conditions to obtain a lysate concentrated enough for subsequent polysome purification, without compromising RNA quality (Figure 2, lanes B–E). The optimal ratio was one volume of egg pellet to four volumes of lysis buffer (1:4 Figure 2, lane E). The extraneous lower bands observed in lanes A, B, C and D correspond to 18S and 28S ribosomal RNA degradation products.

    Two lysis techniques were tested: mechanical shearing through a 25G needle (routinely used for protein lysates in our lab) and Dounce homogenization. In our experiments, the use of a Dounce tissue grinder improved RNA quality (Figure 2, lanes E and F). Using a 7 ml Dounce with the tight B pestle, which leaves a distance of 20–55 μm between pestle and cylinder, we were able to lyse cells without breaking the nucleus to isolate cytoplasmic RNA: the diameter of sea urchin eggs is 100 μm whereas the diameter of the nucleus is less than 20 μm. Proper lysis of the eggs was checked under a light microscope, and 10 strokes were needed to obtain 100% of lysed cells.

    Sea urchin oocytes and embryos contain high nuclease activity that depends on Ca 2+ ions ( 26, 27) a report suggests to rinse them in Ca 2+ -free seawater prior to the lysis step to improve RNA integrity ( 27). In our hands and in the urchin species we used, this step did not significantly improve RNA quality (Figure 2, lanes F–H). However, inclusion of EGTA in the lysis buffer ( 27) improved RNA quality and its reproducibility in the lysates (Figure 2, cf. lanes F, I and J). We also noticed that the high-salt buffer preserved RNA integrity therefore 250 mM KCl was also included in our sea urchin lysis buffer.

    To generate polysomes that accurately reflect the translational status of the cell, ribosome movement on the mRNA must be minimized during sample preparation to effectively prevent ribosomes from running off the mRNA. We therefore added a translation elongation inhibitor. Cycloheximide is the most commonly used inhibitor in polysome profiling protocols described for mammalian cells ( 12). In sea urchin embryos, cycloheximide does not work, however emetine is a very efficient protein synthesis inhibitor ( 28). We therefore included emetine (100 μM) in egg suspensions or embryo cultures 5 min before sample collection, and in the lysis buffer.

    A combination of anti-proteases (aprotinin and leupeptin, for example) or a commercially available anti-protease cocktail is necessary to avoid protein degradation and enhance polysome quality. Additional detergents can be included in the lysate buffer, such as Triton X-100 and sodium deoxycholate to extract membrane-bound polysomes in mammalian tissue or culture cells ( 12, 29) or cetyltrimethylammonium bromide in polysaccharide-containing organisms ( 30). Several RNase inhibitors can be used in combination to fully inhibit RNase activity, such as heparin or vanadyl ribonucleoside complex ( 12, 29). Heparin, known to be an inhibitor of the RT-PCR reaction, can be removed by treating the RNA samples with heparinase I ( 31). In our experiments, we only used a commercial RNase inhibitor (from Promega), with satisfactory output of the purified RNA from sea urchin lysates.

    We recommend carrying out a cytoplasmic RNA integrity check before proceeding with polysome fractionation. RNA was purified from an aliquot of the lysate by acid phenol–chloroform extraction and resolved on an agarose-Tris/Borate/EDTA (TBE) gel to verify that there was no degradation: ribosomal RNA bands appeared as two sharp bands on an agarose gel (see Figure 2, lanes I and J). Quality can be further determined on a Bioanalyzer. The RNA integrity number, which gives a measurement of the degree of degradation, should ideally be close to 10, corresponding to intact RNA ( Supplementary Data ).

    Polysome fractionation

    Sucrose density gradient formation

    Lysates obtained as described above were loaded on a 15–40% linear sucrose density gradient. Two 15 and 40% sucrose buffers were prepared in 10 mM Tris pH 7.4 10 mM MgCl2 250 mM KCl 25 mM EGTA 1 mM DTT. To obtain a linear gradient, we used the Gradient Master device (BioComp) equipped with tubes adapted for a SW41 rotor. We used 6 ml of each sucrose buffer in each gradient tube. In the ultracentrifuge tube, the light sucrose solution was underlayered with the heavy sucrose solution using a cannula attached to a 10 ml syringe. The introduction of air bubbles or mixing of the two solutions should be carefully avoided during this step. The ‘SW41 Short Gradient’ program with the corresponding range of sucrose percentage was run following the manufacturer's instructions. This program lasts a few minutes (2 min 21 s for a 15–40% gradient) and produces up to six reproducible linear gradients in one run. The gradients should be handled with care to avoid disturbance and stored at 4°C for at least 1 h. The percentage of sucrose used for linear gradients can be adjusted to optimize the separation of polysomal fractions according to the biological sample or experiment.

    Sample loading and ultracentrifugation

    When comparing two biological conditions, the same amount of lysate (equivalent ODA260) should be loaded on top of each gradient. The maximum volume of lysate that can be deposited on the gradient is 500 μl, with a maximum of 25 ODA260. In our hands, better separation of the 40S, 60S and ribosome peaks was obtained when 10 ODA260 was loaded on the gradient, typically corresponding to 350 μl of lysate. Loading was carried out by gently pipetting the lysate onto the top of the gradient. Then, the gradient tubes were carefully balanced before starting the ultracentrifugation run at 38000 rpm for 2.5 h in a Beckman SW41 Ti rotor at 4°C. The acceleration was set to its maximum value, and to avoid any disturbance, the deceleration was set to its minimum value. Once the ultracentrifugation run was finished, the gradients were kept at 4°C for immediate fractionation.

    Polysome gradient profile and collection of fractions

    During ultracentrifugation, the Isco Density Gradient Fractionation System was set up according to the manufacturer's recommendations. This density gradient system is equipped with a spectrophotometer with a 254 nm filter (UA-6 UV/VIS detector, Isco), and produces a continuous absorbance profile as the gradient is collected. Fractionation was performed by piercing the bottom of the centrifuged tube to introduce a dense chase solution and raise the intact gradient by bulk flow. Before each experiment, the entire system was washed first with 0.1 M NaOH, then thoroughly with DEPC-treated water. In our experiments, the peristaltic pump was set at a speed corresponding to 0.9 ml/min.

    After centrifugation, the centrifuge tube was connected to a flow cell and the piercing apparatus. The 50% sucrose chase solution was injected by puncturing the tube from the bottom, pushing out the gradient in a continuous manner into the flow cell. Fractions were kept on ice during collection to avoid any RNA degradation. The introduction of air bubbles in the system should be avoided, because bubbles will disturb the gradient. A typical polysome profile (Figure 3A) first showed a peak of A254 absorbing material, containing the untranslated mRNAs, then the two peaks of the small and large ribosomal subunits, the monosome peak and finally the polysomal peaks. Polysome profiles can vary according to the general translation activity. For example in unfertilized sea urchin eggs, <2% of the ribosomes are engaged in polysomes and polysome peaks are barely detectable (( 19) Figure 3A). After fertilization, protein synthesis activity increases and the proportion of ribosomes in polysomes increases as development proceeds (Figure 3A).

    (A) Polysome gradient profile during early development. Optical density profiles (ODA254) of polysome gradient profiles are shown, corresponding to unfertilized eggs (UnF), 1 h post-fertilization embryos (F) and late-blastula stage embryos 30 h post-fertilization (Blastula). The areas under the curve (AUC) of polysomes and monosomes were measured, and the polysome:monosome ratio was then calculated for the three developmental stages error bars represent SEM and statistics were done using Student's t-test (P-value < 0.01). (B–E) Polysome gradient profiles and corresponding RNA profiles after treatment with polysome disrupters. Optical density profiles and extracted RNAs from polysome gradients of fertilized eggs (B), treated with 30 mM EDTA (C), 2 mM puromycin in vitro (D) and 0.6 mM puromycin in vivo (E) are shown. The RNAs from each fractions of polysome gradient were separated on 2% agarose-TBE gels.

    (A) Polysome gradient profile during early development. Optical density profiles (ODA254) of polysome gradient profiles are shown, corresponding to unfertilized eggs (UnF), 1 h post-fertilization embryos (F) and late-blastula stage embryos 30 h post-fertilization (Blastula). The areas under the curve (AUC) of polysomes and monosomes were measured, and the polysome:monosome ratio was then calculated for the three developmental stages error bars represent SEM and statistics were done using Student's t-test (P-value < 0.01). (B–E) Polysome gradient profiles and corresponding RNA profiles after treatment with polysome disrupters. Optical density profiles and extracted RNAs from polysome gradients of fertilized eggs (B), treated with 30 mM EDTA (C), 2 mM puromycin in vitro (D) and 0.6 mM puromycin in vivo (E) are shown. The RNAs from each fractions of polysome gradient were separated on 2% agarose-TBE gels.

    RNA purification and profiles

    The RNAs of each fraction were extracted with one volume of acid phenol–chloroform (vol:vol), and precipitated with one volume of isopropanol. Ethanol–sodium acetate precipitation cannot be used, due to the high sucrose concentration in the last fractions. RNAs were pelleted in a tabletop centrifuge at 13000 rpm for 10 min at 4°C, washed with 70% ethanol, pelleted again and air-dried for 20 min. RNAs from each fraction were resuspended in the same volume of RNase-free water (one-twentieth of the original fraction volume). An aliquot of each fraction was used to check RNA quality on an agarose gel. The polysome profile of fertilized embryos and the corresponding RNA profile of the 15–40% sucrose gradient separated on an agarose gel are shown in Figure 3B. In the first fractions, only low molecular weight RNAs were visible, then fractions containing only 40S subunits were isolated as shown by the presence of the 18S rRNA band, followed by the 60S peak, as shown by the 28S rRNA band. In the middle of the gradient, the bands were more intense, due to the high proportion of monosomes. Starting from fraction 13, 28S and 18S RNAs were present with a 28S:18S ratio equal to 2:1, representing polysomal fractions. On average, starting from 10 ODA260 loaded on the gradient, typical RNA yields were 750 ng per polysome fraction.

    Protocols often recommend using proteinase K treatment before RNA extraction (10 mM Tris pH 8, 1 mM ethylenediaminetetraacetic acid (EDTA), 0.5% sodium dodecyl sulphate and 200 μg/ml proteinase K, for 25 min at 50°C) to enhance recovery of RNAs from polysome gradients ( 32). Although it may help for specific samples, in our hands, the proteinase K treatment did not significantly improve RNA quality or quantity (data not shown).

    Analysis of the translational status of mRNAs

    A critical point of translation analyses is to verify that mRNAs present in polysome fractions are associated with translating polysomes and do not merely co-sediment with them. In the cell, untranslated mRNAs are associated with RNA-binding proteins in so-called messenger ribonucleoparticles (mRNPs) or with stalled polysomes, which may sediment in the same fractions as polysomes when purified on sucrose gradients ( 19, 33). To distinguish between active polysomes and co-sedimenting mRNPs or stalled polysomes, we treated our samples with a polysome disrupter prior to polysome purification. The two commonly used disrupters are EDTA and puromycin. EDTA chelates Mg 2+ ions and dissociates the subunits of the ribosome. EDTA (30 mM final) was added after lysate preparation, prior to gradient loading. Gradients should also be supplemented with 10 mM EDTA and prepared without Mg 2+ . Given that EDTA can potentially affect mRNPs, whose formation may depend on Mg 2+ , the antibiotic puromycin is usually preferred. Puromycin causes the dissociation of the ribosomes in the elongation step of translation, and only affects active polysomes ( 34). Puromycin can be added in vivo prior to lysis (0.6 mM in sea urchin egg suspension or embryo culture) or in vitro after lysate preparation (2 mM in lysate), when samples are not easily available for puromycin in vivo incubation ( 23, 35). In both cases, the lysate is supplemented with 500 mM KCl, incubated for 15 min at 4°C then for 15 min at 37°C before loading on the sucrose gradient ( 35).

    EDTA treatment disrupted the gradient profile, with a high loss of RNA integrity (Figure 3C). Adding more RNase inhibitor to lysates and in the sucrose gradient improved RNA quality (data not shown). In contrast, the use of puromycin either in vitro or in vivo did not affect RNA quality after purification. RNAs from polysome fractions shifted toward lighter fractions in puromycin-treated lysates (Figure 3D and E). We therefore used the puromycin treatment for assessing translation of a specific mRNA.

    Following polysome profiling and using the puromycin controls in parallel, the translational status of a specific mRNA can be assessed by Northern blot or RT-PCR. The presence of the mRNA can be detected in each fraction, and after quantification of the signal, specific mRNA distribution was expressed as a percentage of the total signal. An illustration of this approach is given below, by analyzing the translational status of mRNAs before and after fertilization in sea urchin.


    Discussion

    Here, we investigate the translational regulation of APC-dependent RNAs which are targeted to cell protrusions. We find that the cytoplasmic position of APC-dependent RNAs does not affect their translation, since they can be translated similarly in both internal and peripheral locations. Rather, translation of APC-dependent RNAs is coordinated with specific peripheral cellular processes, being activated at extending protrusions/lamellipodia and suppressed upon protrusion retraction. Silencing is coupled to a change in the physical state of the RNAs manifested as single RNAs clustering into heterogenous granules at the tips of protrusions.

    This mode of regulation is distinct from the one proposed for several other localized transcripts, whereby RNAs are transported in a silenced state and are translationally activated only upon reaching the final destination or upon receipt of specific signals (Besse and Ephrussi, 2008 Buxbaum et al., 2015 Jung et al., 2014). A reason for this latter type of regulation has been proposed to be the need to prevent protein appearance at sites, or times, where it might be deleterious. Indeed, premature appearance of the transcriptional repressor Ash1p in the mother cell during budding suppresses transcriptional programs in both mother and daughter cells and prevents mating-type switching (Long et al., 1997). Similarly, disrupting the timing of translational activation within neuronal axons or dendrites can lead to aberrant axonal pathfinding or synaptic responses (Colak et al., 2013 Holt and Schuman, 2013 Jung et al., 2012).

    Our observation of translation regardless of cytoplasmic location suggests that the proteins encoded by APC-dependent RNAs can be produced in internal regions without deleterious effects. We propose that this mode of regulation could additionally have functional implications for the encoded proteins. Specifically, translation in local environments can attribute proteins with different properties. This could result from differential protein modifications or through proximity to protein partners, which during co-translational assembly could affect the type or efficiency of multimeric complex formation (Basu et al., 2011 Jung et al., 2014 Shiber et al., 2018 Shieh et al., 2015). In light of these ideas, we suggest that a single mRNA, as it is being continuously translated at various stages during its transport to the periphery, could give rise to protein copies which have different properties, and therefore functional potential, depending on the local micro-environment they are translated into.

    We additionally show that translation of APC-dependent RNAs is specifically suppressed in retracting protrusions and this silencing is associated with the formation of multimeric heterogeneous clusters. Formation of these clusters is reduced by treatments that prevent RNA transport to the periphery and is increased upon global translational inhibition, suggesting that silencing is a limiting step in their formation. Nevertheless, silencing can occur outside of clusters, since a large proportion of RNAs are observed as single particles in retracting, translationally-silent protrusions. The appearance of these heterogenous RNA clusters at the tips of protrusions, and their relation to translation, are reminiscent of other types of RNA granules formed by liquid-liquid phase separation (Courchaine et al., 2016 Weber and Brangwynne, 2012). For example, stress granules (SGs) and processing bodies (PBs) are sites of dynamic concentration of RNA molecules, but in both cases translational silencing or decay can occur regardless of RNA localization within granules (Halstead et al., 2015 Panas et al., 2016 Perez-Pepe et al., 2018). SGs and PBs form throughout the cytoplasm under normal conditions or in response to stress. An interesting distinction of the clusters described here is that their formation is induced in particular subcellular regions associated with protrusion retraction, suggesting that their assembly/disassembly is controlled by spatial signals.

    The silencing of Rab13, and likely of other APC-dependent RNAs, at retracting protrusions is contrasted by their translational activation in extending lamellipodia. Given that localization of APC-dependent RNAs to the periphery is important for cell migration (Wang et al., 2017) these data could point to a functional role for spatially segregating translation such that local protein production occurs in actively extending regions, while being suppressed in retracting areas. This would imply the existence of a dynamic regulatory mechanism that coordinates APC-dependent RNA translation with the continuous protrusion and retraction cycles that characterize cellular migration (Ji et al., 2008 Tkachenko et al., 2011).

    A potential underlying mechanism for local silencing and granule formation could rely on spatially restricted phosphorylation/dephosphorylation events, which have been shown to affect the propensity of RNA-binding proteins to form phase-separated granules or to bind to RNAs (Monahan et al., 2017 Murray et al., 2017 Thapar, 2015). In this regard, it is interesting that proteins associating with APC-dependent RNAs include the translational regulator FMRP and the RNA-binding protein FUS, one of the paradigm proteins used in understanding phase transitions (Mili et al., 2008 Yasuda et al., 2017 Yasuda et al., 2013). Local modifications of FMRP or FUS could underlie the observed regulation. Additional local events, including local maturation of miRNAs, could be envisioned (Sambandan et al., 2017).

    With regards to the assembly/disassembly dynamics of peripheral clusters and the fate of the sequestered RNAs, limitations in the duration of live-imaging we can accomplish do not permit concrete conclusions. However, our current data indicate that the extent of cluster formation might be influenced by the rate of retraction. Specifically, in NIH/3T3 cells, the majority of peripheral clusters are characterized by a visible accumulation of polyA signal, indicative of a high concentration of heterogeneous RNA species. By contrast, a substantial fraction of retracting MDA-MB-231 protrusions do not exhibit obvious polyA accumulation (Figure 9). A prominent difference between the two cell types is the speed with which protrusions retract. Contractile protrusions of NIH/3T3 cells can persist for a long time, while MDA-MB-231 protrusions quickly retract within a few minutes (compare Videos 11 and 12 to Video 13). While we cannot rule out other interpretations, such as the existence of deadenylated transcripts, we favor the idea that slowly retracting protrusions allow for the build-up of large heterogenous granules, while in faster retracting protrusions peripheral granules are transient, and rapidly disassemble either through RNA degradation or release into the cytosol.

    The existence of translationally silent multimeric RNA clusters also offers a potential explanation for the slight decrease of APC-dependent RNAs found in the heavy RNP fraction under conditions that disrupt transport to the periphery (Figures 1D and 2B). Translationally silent, higher-order RNP complexes can sediment at sucrose density gradient fractions heavier than their translated counterparts (Chekulaeva et al., 2006). By analogy, the peripheral clusters of APC-dependent RNAs likely account for some of the RNA found at heavier fractions of the gradient. Reduction of their formation upon parthenolide treatment (Figure 10) could likely account for the apparent shift of APC-dependent RNAs towards the lighter polysome fraction of the gradient (Figures 1D and 2B).

    We had previously reported that APC-dependent RNAs can also be found in internal cytoplasmic granules induced by expression of FUS mutants carrying ALS-associated mutations. Intriguingly, in these FUS-granules APC-dependent RNAs are translationally active (Yasuda et al., 2013). Taken together, these observations raise the possibility that, at least for APC-dependent RNAs, their physical partitioning into granules is not the sole determinant, but acts in combination with the particular local environment to determine the eventual impact on their translation status. It would be interesting to further investigate how the composition of peripheral polyA granules differs from other internal cytoplasmic RNA granules with regards to both RNA and protein constituents.

    We note that the above study describes the regulation of APC-dependent RNAs in migrating mesenchymal cells. It would be interesting to explore how translation and transport of this RNA group is carried out in larger and more stably polarized cells such as neurons.

    Overall, we describe here a distinct mode of translational regulation of localized RNAs. These findings provide a different perspective towards understanding how local translation can influence protein activities and how these regulatory mechanisms could be integrated with dynamic cellular behaviors.


    Results

    Very few ribosome footprints on unspliced HAC1 mRNA

    We recently reported improved ribosome-footprint profiles (Ingolia et al., 2009) and mRNA-abundance measurements from exponentially growing S. cerevisiae (Weinberg et al., 2016). Under these growth conditions HAC1 mRNA is almost entirely unspliced (Figure 1—figure supplement 1A) and is distributed across a sucrose gradient with

    50% in the non-translating fractions and the remaining

    50% extending across all of the translating (i.e., 80S and larger) fractions without substantial enrichment in any particular fraction (Figure 1A), similar to previous observations (Arava et al., 2003 Chapman and Walter, 1997 Cox and Walter, 1996 Kuhn et al., 2001 Mori et al., 2010 Park et al., 2011 Payne et al., 2008 Rüegsegger et al., 2001 Sathe et al., 2015). In contrast, the well-translated actin (ACT1) mRNA is essentially absent from the non-translating fractions and is found mostly in large polysomes. Based on the sedimentation of HAC1 mRNPs, we predicted that the mRNA would generate a large number of ribosome-protected fragments, in a quantity only

    2-fold fewer than similarly abundant mRNAs (based on the fraction of HAC1 mRNA in the untranslated fractions). Strikingly, however, after normalizing for mRNA abundance HAC1 generates the fewest ribosome-protected fragments among all expressed yeast genes (Figure 1B)—and

    50-fold fewer than expected from the polysome profile. Rather than providing evidence for stalled ribosomes on HAC1 u mRNA, instead these observations suggest that either ribosomes are stalled on the mRNA in a closely packed configuration that prevents nuclease cleavage between ribosomes, which would eliminate the

    28-nucleotide fragments that are sequenced in the ribosome-profiling method (Figure 1C, middle) or that there are not stalled ribosomes on HAC1 u mRNA (Figure 1C, bottom).

    Ribosome density on unspliced HAC1 mRNA.

    (A) Polysome analysis of HAC1 and ACT1 mRNAs. Extracts prepared from exponentially growing yeast cells were fractionated on 10–50% sucrose gradients, with absorbance at 260 nm monitored (top). The relative distributions of HAC1 and ACT1 mRNAs across fractions were determined by qRT-PCR (bottom). Shown are the mean ± SEM with n = 2 (i.e., the range), expressed as a fraction of the total mRNA detected. (B) Histogram of ribosome densities measured by ribosome profiling and RNA-seq. The ratio of the number of ribosome-protected fragments (RPFs) to the number of RNA-seq reads (mRNA counts) was calculated for each of 4838 expressed yeast genes (data from Weinberg et al., 2016). Shown is the distribution of log-transformed ratios in bins of 0.05, with the position of HAC1 indicated. (C) Possible scenarios to explain a lack of RPFs. While an mRNA with average ribosome density will generate many

    28 nucleotide (nt) RPFs (top), the close packing of stacked ribosomes could inhibit the RNase digestion between ribosomes required to generate

    28 nt RPFs (middle). Alternatively, an mRNA that does not contain translating ribosomes would not generate RPFs (bottom). (D) Polysome analysis of HAC1 mRNA variants with shortened ORFs. G-to-T point mutations were introduced into the first exon of HAC1 to generate premature stop codons, with the resulting ORFs shown as thick colored boxes (constitutive 5′- and 3′-UTRs located within exons 1 and 2, respectively, are shown as thin black lines other untranslated regions are depicted as thin colored boxes and the coding regions of exons 1 (teal) and 2 (purple) are labeled as 'HAC1 exon1' and ‘exon2’, respectively). The maximum number of ribosomes that could be accommodated was calculated based on each ribosome occupying 28 nt. Polysome analysis was performed as in (A), with data for wild-type HAC1 from (A) duplicated for comparison. (E) Effects of heparin on polysome analysis. Purified uncapped luciferase (luc) RNA was added to either lysate or lysis buffer in the absence (–) or presence (+) of 0.2 mg/ml heparin. Polysome analysis was performed as in (A) with absorbance at 260 nm monitored (top), and the relative distributions of exogenous luc RNA (middle) and endogenous HAC1 and ACT1 mRNAs (bottom in lysate only) were determined. (F) Refined polysome analysis of HAC1 mRNAs. Extracts were prepared in heparin-containing lysis buffer from strains shown in (D). Polysome analysis was performed as in (A). (G) Polysome analysis of HAC1 mRNAs during the UPR. Strains shown in (D) were grown to mid-log phase and treated with 8 mM DTT for 20 min before harvesting. Extracts were prepared in heparin-containing lysis buffer, and polysome analysis was performed as in (A). Dotted lines indicate the fractions after which the corresponding color-coded mutant mRNAs would not be expected to sediment based on ORF length.

    Inhibited translation initiation revealed by polysome analyses

    Although the polysome-like sedimentation of HAC1 u mRNA indicates ribosome association, it does not reveal if the ribosomes associated with the mRNA were ever engaged in its translation. Alternatively, the associated ribosomes might be bound in a conformation that is unrelated to translation of HAC1 u mRNA. We therefore devised an experiment to definitively determine whether the ribosomes bound to HAC1 u mRNA reflect ribosomes that were engaged in its translation. To do so, we took advantage of the observation that HAC1 mRNA is distributed across all of the translating fractions of a sucrose gradient (Figure 1A). If the deep sedimentation is due to multiple translating ribosomes being stalled on a single mRNA, then reducing the number of translating ribosomes that can fit on the mRNA should shift the sedimentation pattern toward lighter fractions. We designed a series of constructs containing point mutations in the first exon of HAC1 that created premature termination codons, which reduce the size of the ORF and thereby limit the number of translating ribosomes (Figure 1D, top). To ensure that the mutant alleles were expressed at near wild-type levels, we replaced the endogenous HAC1 allele without disrupting flanking regulatory regions. Remarkably, each of the mutant mRNAs had a sedimentation pattern that was indistinguishable from the wild-type mRNA (Figure 1D, bottom). In the most extreme case, the mRNA containing a 21-nucleotide ORF that can only accommodate a single translating ribosome still co-sedimented with polysomes containing upwards of 10 ribosomes (fraction 14 of the gradient). These data provide direct evidence that the polysome-like sedimentation of HAC1 u mRNA is not due to stalled ribosomes on the mRNA.

    We hypothesized that the ribosome association of HAC1 u mRNA was instead due to non-specific interactions between the mRNA and bona fide polysomes formed on other mRNAs. To evaluate the extent of such non-specific interactions, we introduced an exogenous control RNA that should not be translated: an uncapped luciferase-encoding RNA purified from an in vitro transcription reaction. We analyzed the sedimentation behavior of this control RNA when it was added to lysis buffer compared to when it was added to the yeast lysate prior to centrifugation. Surprisingly, some of the exogenous RNA was found in the translating fractions of the lysate—a behavior not observed in lysis buffer alone (Figure 1E, middle). Through extensive optimization, we found that the addition of heparin to the lysis buffer (at a concentration of 0.2 mg/ml) was sufficient to largely prevent the deep sedimentation of exogenous RNA. The addition of heparin also had a major effect on the sedimentation of HAC1 mRNA, as most (83%) now sedimented in the non-translating fractions of the gradient (Figure 1E, bottom). In contrast, the sedimentation of ACT1 mRNA (Figure 1E, bottom) and the overall polysome profile (Figure 1E, top) were largely unchanged, suggesting that heparin competed away non-specific interactions without disrupting bona fide polysomes.

    Based on these results, we re-analyzed the HAC1 mutants with shortened ORFs using heparin-containing lysis buffer. Under these conditions, the polysome co-sedimentation of each of the mRNAs was greatly reduced (from

    50% to < 20%) but there was still no difference among the constructs (Figure 1F), providing further evidence against stalled ribosomes on HAC1 u mRNA. Importantly, when we treated cells with the reducing agent dithiothreitol (DTT) to induce the UPR and concomitant splicing of HAC1 mRNA (Figure 1—figure supplement 1C) and repeated the experiment, the variant mRNAs now displayed the expected differential sedimentation based on ORF length (Figure 1G), validating our experimental design. Together, these results demonstrate that at steady state HAC1 u mRNA is not associated with either actively elongating or stalled ribosomes. This indicates that the primary block to production of Hac1 u p is at the stage of translation initiation, not translation elongation as previously proposed (Chapman and Walter, 1997 Richter and Coller, 2015 Rüegsegger et al., 2001).

    An additional silencing mechanism downstream of translation initiation

    The absence of ribosomes on HAC1 u mRNA suggests that translation initiation is inhibited. To further understand the mechanism of this inhibition, we used a green fluorescent protein (GFP) reporter system that has been previously shown to recapitulate post-transcriptional silencing by the HAC1 intron (Chapman and Walter, 1997 Rüegsegger et al., 2001). We replaced the first exon of HAC1 with the GFP ORF lacking its own stop codon (Figure 2B), which allowed us to quantitatively analyze post-transcriptional silencing of GFP using a combination of flow cytometry (protein abundance), quantitative RT-PCR (mRNA abundance), and sucrose gradient fractionation (ribosome density). When GFP was embedded in an otherwise wild-type HAC1 context the mRNA sedimented almost entirely in the non-translating fractions (Figure 2A), and there was no detectable fluorescence above background (Figure 2C). In contrast, cells expressing a reporter construct missing the entire intron displayed a strong fluorescence signal (Figure 2C), and most of the mRNA was found in the translating fractions (Figure 2A). Thus, the reporter GFP mRNA behaves similarly to endogenous HAC1 mRNA.

    Contribution of long-range base pairing to intron-dependent silencing.

    (A) Polysome analysis of reporter mRNAs. Extracts were prepared in heparin-containing lysis buffer from strains expressing the GFP reporter mRNAs depicted in (B). Polysome analysis was performed as in Figure 1A, with data for wild-type HAC1 from Figure 1F duplicated for comparison. (B) Design of reporter mRNAs. Constructs are depicted as in Figure 1D, with the dotted line indicating a deleted region. Colored stars indicate mutations to the base-pairing region, with specific nucleotide changes shown below in red. (C) Flow cytometry analysis of reporter strains. Strains expressing the GFP reporter mRNAs depicted in (B) were grown to mid-log phase and analyzed by flow cytometry. Plotted is the median GFP intensity (normalized to cell size) of the cell population relative to background fluorescence in the wild-type (no GFP) strain with error bars indicating quartiles of the cell population, all averaged across replicates (n = 2–7).

    To determine whether 5′-UTR–intron base pairing is required to prevent ribosome loading on the reporter mRNA, we designed constructs in which the sequence implicated in base-pairing interactions in either the 5′-UTR or the intron was mutated to its complement to disrupt the interaction (Figure 2B, bottom), as was done previously (Rüegsegger et al., 2001). Both mutant mRNAs sedimented mostly in the translating fractions in a manner that was similar to the intronless construct (Figure 2A). When we combined the 5′-UTR and intron mutations and thereby restored base pairing, the mRNA was now found almost exclusively in the non-translating fractions and resembled the original reporter mRNA. These results demonstrate that base pairing between the 5′-UTR and intron is required to prevent ribosome loading by directly impeding the binding or progress of the scanning ribosome.

    Given that the base-pairing mutant mRNAs were loaded with ribosomes similarly to the intronless mRNA (Figure 2A), we expected to observe GFP expression from the mutant mRNAs that was similar to that of the intronless construct. Remarkably, however, neither of the strains expressing a base-pairing mutant mRNA had any GFP detectable by either flow cytometry (Figure 2C) or immunoblotting (Figure 2—figure supplement 1). The lack of GFP signal despite polysome sedimentation was not due to low mRNA abundance, as the mutant mRNAs were present at similar levels compared to the intronless mRNA (Figure 2—figure supplement 1A, compare construct 3 with constructs 5–6). These results suggest that an additional silencing mechanism acting downstream of translation initiation prevents GFP accumulation when base pairing is disrupted.

    Post-translational silencing by the intron-encoded C-terminal tail

    In addition to removing the intron portion of the base-pairing interaction, splicing of HAC1 mRNA also alters the C-terminal tail of the encoded protein: The ORF in HAC1 u mRNA has a 10-amino-acid tail encoded by the intron, which in HAC1 i mRNA is replaced by an 18-amino-acid tail encoded by the second exon (Figure 3A). The fact that the polypeptide tails encoded by the HAC1 u and HAC1 i mRNAs are different suggests that the 10-amino-acid tail unique to Hac1 u p may be functionally important. We therefore hypothesized that the intron-encoded C-terminal tail may be involved in the additional silencing mechanism revealed by our base-pairing mutant reporters (Figure 2).

    Post-translational silencing mediated by the intron-encoded C-terminal tail.

    (A) Schematic of HAC1 mRNA splicing. The proteins encoded by HAC1 u and HAC1 i mRNAs differ in their C-terminal tails, with the amino acid sequences indicated. (B) Design of reporter mRNAs. Black shading indicates recoding, with the original and recoded sequences depicted below (mutations in red). Untranslated regions colored red correspond to those of the TMA7 mRNA, with the reporter gene integrated at the TMA7 rather than HAC1 locus. Otherwise constructs are depicted as in Figure 2B. (C) Flow cytometry analysis of reporter strains. Strains expressing the GFP reporter mRNAs depicted in (B) were analyzed as in Figure 2C, with data for the first three strains duplicated from Figure 2B for comparison. (D) Polysome analysis of reporter mRNAs. Extracts were prepared in heparin-containing lysis buffer from strains expressing the GFP reporter mRNAs indicated in (B). Polysome analysis was performed as in Figure 1A, with data for the wild-type and intronless GFP reporters from Figure 2A duplicated for comparison. (E) Differentiating between co-translational and post-translational silencing mechanisms. Left: Schematic of reporter construct that generates two separate polypeptides from each round of translation. Right: Extracts were prepared from strains expressing reporter mRNAs that either encoded the 10-amino-acid C-terminal tail of Hac1 u p (+tail) or contained a stop codon just before the tail (–tail). Immunoblotting was used to detect HA-tagged mRuby (top) and GFP (bottom), with actin as a loading control. Two biological replicates are shown for each genotype.

    To investigate this possibility, we designed a reporter construct in which we removed the entire intron except for the 5′ end that codes for the 10-amino-acid tail of Hac1 u p (Figure 3B, fourth construct). Remarkably, cells expressing this construct had no detectable GFP (Figure 3C), despite the corresponding mRNA being abundant (Figure 3—figure supplement 1A) and detected on polysomes due to the absence of the base-pairing interaction (Figure 3D). Thus, the coding region at the 5′ end of the intron is sufficient for complete silencing of the GFP reporter. Introducing a termination codon between GFP and the intron restored robust fluorescence comparable to that observed for the intronless construct, which implicated translation of the 5′ end of the intron as required for silencing. Furthermore, making 10 nucleotide changes that maintained the coding potential of the intron-encoded tail (Figure 3B, bottom) had no effect on silencing (Figure 3C), suggesting that the amino-acid sequence encoded by the 5′ end of the intron is more important than the nucleotide sequence itself.

    To determine whether the 10-amino-acid element alone was sufficient for silencing in the absence of any other HAC1 sequences, we expressed GFP from a different locus in the yeast genome (TMA7) either with or without the 10-amino-acid tail. When the 10-amino-acid sequence was either absent or not translated due to a premature stop codon, we observed robust GFP signal (Figure 3C). In contrast, there was no fluorescence detected in cells expressing GFP containing the 10-amino-acid tail. Thus, the C-terminal tail of Hac1 u p is sufficient for silencing independently of the rest of the HAC1 intron and any other HAC1 sequences.

    Having established the effects of the 10-amino-acid tail in isolation, we next examined the effects of the tail when translation initiation was inhibited by the base-pairing interaction. In an otherwise wild-type HAC1 context, preventing translation of the 10-amino-acid tail either by deleting the entire sequence or by introducing a stop codon caused GFP to accumulate to low but detectable levels (Figure 3C) without greatly affecting the sedimentation of the corresponding mRNAs (Figure 3D). Thus, even when base pairing is intact there is some low-level accumulation of GFP that is normally suppressed by the 10-amino-acid silencing element.

    In the context of reporter constructs lacking the intron-encoded C-terminal tail, mutations in either the 5′-UTR or intron that disrupted base pairing greatly increased the amount of GFP, while the compensatory double-mutant construct with restored base pairing had only low levels of GFP (Figure 3C and Figure 3—figure supplement 1B). These results are in agreement with previous GFP reporter experiments, which used constructs that contained a stop codon between the GFP and intron sequences and therefore had inadvertently eliminated the effects of the 10-amino-acid tail (Chapman and Walter, 1997 Rüegsegger et al., 2001).

    Together, our GFP reporter experiments indicate that the HAC1 intron mediates post-transcriptional silencing through a pair of independent but partially redundant mechanisms: base-pairing interactions with the 5′-UTR that inhibit translation initiation, and a novel silencing mechanism mediated by the intron-encoded C-terminal tail of Hac1 u p. Robust expression requires that both silencing mechanisms be inactivated, as would happen simultaneously when the intron is removed by Ire1p-dependent splicing.

    What is the mechanism by which the intron-encoded tail of Hac1 u p silences gene expression? Because disrupting translation of the tail elevated protein levels (Figure 3C) without affecting polysome formation (Figure 3D), we inferred that the tail was exerting its effect downstream of translation initiation. We therefore reasoned that the 10-amino-acid sequence was acting either by halting translation across the entire mRNA (after polysome formation) or by promoting protein degradation. Our inability to detect GFP containing the 10-amino-acid tail (Figure 3C) prevented us from directly comparing the half lives of GFP with and without the tail. Instead, to distinguish between stalled elongation and protein degradation, we designed a reporter construct that generates two separate polypeptides from a single round of translation through co-translational ‘cleavage’ mediated by a viral 2A peptide (Sharma et al., 2012). After screening for a 2A peptide sequence that functions efficiently in S. cerevisiae (Figure 3—figure supplement 2), we designed a construct containing HA-mRuby and GFP sequences separated by the P2A peptide (derived from porcine teschovirus-1). The GFP sequence downstream of P2A was appended with the 10-amino-acid tail (or was not, as a control), which should lead to the absence of detectable GFP regardless of the mechanism of action. In contrast, the upstream HA-tagged mRuby should only be detected if translation itself is not affected by the 10-amino-acid sequence, since it reports on the number of rounds of translation but is not covalently linked to the inhibitory tail. Assaying for GFP by immunoblotting revealed that accumulation of the protein was suppressed by the 10-amino-acid tail, as expected (Figure 3E). In contrast, HA-mRuby was detected at similar levels whether or not the tail was included in the construct. These results indicate that the tail functions downstream of translation, likely by acting as a ‘degron’ (Varshavsky, 1991) that targets the protein for immediate degradation after synthesis (Cox and Walter, 1996).

    Identification of DUH1 through a genetic selection

    If the C-terminal tail of Hac1 u p functions as a degron, additional proteins may be involved in recognizing the degron and targeting the covalently linked protein for degradation. To identify such trans-acting factors, we used a genetic approach that took advantage of the strong silencing phenotype imparted by the 10-amino-acid tail alone (Figure 3C). We constructed strains in which the first exon of HAC1 was replaced by HIS3, which we reasoned might behave like the analogous GFP reporter genes and be completely silenced by the 10-amino-acid sequence. Similarly to GFP, replacing the first exon of HAC1 with HIS3 but keeping the HAC1 locus otherwise intact prevented expression of His3p as evidenced by histidine auxotrophy (Figure 4A). Removing the entire intron restored His3p expression and growth of the corresponding strain on medium lacking histidine, suggesting that silencing was mediated by the HAC1 intron. Strains containing a stop codon between HIS3 and the intron (to prevent translation of the 10-amino-acid degron) had a low but detectable level of growth on histidine-lacking medium (Figure 4A), consistent with the weak fluorescence signal observed from the corresponding GFP reporter construct (Figure 3C). On the other hand, strains containing His3p appended with the C-terminal tail of Hac1 u p but no other elements of the HAC1 intron could not grow on medium lacking histidine. These results indicate that the degron is sufficient for functional silencing of HIS3 expression, providing a useful genetic tool to identify additional genes involved in the degron-dependent silencing mechanism.

    Identification of DUH1 through a genetic selection.

    (A) Evaluating a genetic reporter for intron-dependent silencing. Strains expressing the indicated HAC1-based HIS3 reporter mRNAs (depicted as in Figure 2B) either without (–) or with (+) an N-terminal HA tag were grown to saturation, and 10-fold dilution series were plated on either SC or SC–His media. (B) Flowchart of genetic selection for dds mutants. After selecting for spontaneous mutants that could grow on medium lacking histidine, restreaked clones were filtered out for cis mutants, verified to be expressing HA-His3p, and a subset analyzed by whole-genome sequencing. (C) Chromosome map of mutations identified by whole-genome sequencing. Each color corresponds to a different dds mutant strain. Shown on the right is the DUH1 locus (YJL149W), with locations of nucleotide changes (with respect to DUH1 start codon) and corresponding amino changes listed below. (D) Effect of DUH1 disruption on expression of the genetic reporter. Strains expressing the indicated reporter mRNAs depicted in (A) in either a DUH1 or duh1∆ (two independent clones) background were grown to mid-log phase. Extracts were prepared and immunoblotted for HA-His3p and actin loading control. (E) Flow cytometry analysis of reporter strains. Strains expressing the indicated GFP reporter mRNAs in either a DUH1 or duh1∆ background were analyzed as in Figure 2C. Data is plotted as in Figure 2C (n = 2 for all strains).

    Although we had initially intended to use chemical mutagenesis to generate silencing-defective mutants, upon streaking out the selection strain on histidine-lacking medium we noticed that a small number of slow-growing colonies appeared even without mutagen treatment. We reasoned that such spontaneous suppressor strains would have very few mutations, making it possible to identify suppressor mutations by whole-genome sequencing without requiring backcrossing or forming complementation groups. Thus, to isolate mutants with defective d egron- d ependent s ilencing ('dds mutants') we simply plated the selection strain expressing HA-tagged His3p with the 10-amino-acid tail on histidine-lacking medium and isolated the rare single colonies that grew for further analysis (Figure 4B). From five independent platings we isolated a total of 123 mutant strains that, when re-streaked, could grow on medium lacking histidine. Sanger sequencing of the C-terminal region of the reporter gene in each mutant identified 15 strains harboring a cis mutation that either altered the sequence of the degron, introduced a premature stop codon before or within the degron, or removed the stop codon of the degron resulting in a six-amino-acid C-terminal extension (Figure 4—figure supplement 1A)—all of which provided confirmation that the genetic selection worked as desired.

    From the strains that displayed histidine prototrophy, we picked 35 (including four cis mutants) to analyze by immunoblotting and found that all had detectable HA-His3p (Figure 4—figure supplement 1B). We selected 20 of these strains with unknown mutations for whole-genome sequencing (as well as the parental selection strain as a reference, and three strains with known mutations in the degron as positive controls for our variant-calling procedure), taking care to select strains that varied widely in HA-His3p abundance or growth rate to minimize our chances of sequencing the same mutation in multiple strains. We sequenced the 24 genomes together in a single lane of a HiSeq sequencer using 50-nucleotide single-end reads, which provided 38–74X coverage (9.3–18.0 million reads) of each yeast genome. We then used standard mapping and variant-calling tools (BWA and FreeBayes, respectively) to identify variants that were absent from the parental genome, which successfully recovered the positive-control cis mutants. Remarkably, 17 out of the 20 strains that we sequenced contained a mutation in the same gene YJL149W (Figure 4C and Figure 4—figure supplement 1C), and in each case we confirmed the mutation by Sanger sequencing (Figure 4—figure supplement 1D). Because no other ORF was found mutated in more than one of the 20 mutant strains (Figure 4—figure supplement 1C), we focused our follow-up efforts on YJL149W.

    YJL149W had previously been named DAS1 for ' D st1-delta 6- A zauracil S ensitivity 1 ' when it was identified in a genetic screen unrelated to the UPR (Gómez-Herreros et al., 2012). Based on the protein’s domain structure (containing an F-box domain and leucine-rich repeats) and physical interactions with the SCF core components Cdc53p and Skp1p (Willems et al., 1999), YJL149W was annotated as a 'putative SCF ubiquitin ligase F-box protein' (Cherry et al., 2012). F-box proteins act as adapters to target substrates for ubiquitination and subsequent degradation by the proteasome (Skaar et al., 2013). We therefore propose to rename this gene DUH1 for ' D egrader of U nspliced H AC1 gene product 1 ' to reflect its role in the degradation of Hac1 u p, as we demonstrate later.

    The DUH1 variants that we identified in our 17 strains comprised 16 different mutations, of which 8 were nonsense, 7 were missense, and 1 was a frameshift (Figure 4C, right). The nonsense mutations tended to cluster in the first half of the ORF, suggesting that they likely functioned as null alleles. In addition, three of the 17 strains containing mutations in DUH1 did not contain any other mutations, implicating the DUH1 mutations as causative for the phenotype. We therefore tested whether knocking-out DUH1 (which is non-essential) in the original selection strain recapitulated the de-silencing phenotype. Disrupting DUH1 led to a dramatic increase in the steady-state abundance of HA-His3p containing the 10-amino-acid tail, while having no effect on HA-His3p lacking the tail or constructs repressed by long-range base pairing (Figure 4D). Analogously, deleting DUH1 in the GFP reporter strains completely eliminated the silencing effect of the Hac1 u p tail but had no effect on base-pairing-mediated silencing (Figure 4E and Figure 4—figure supplement 2). Notably, in the absence of DUH1 the GFP reporter in a wild-type HAC1 context was now expressed at the same leaky level as previously seen for the reporter in which a stop codon was positioned between GFP and the intron, indicating that degron-dependent silencing is required to suppress leaky GFP expression. Together, these results provide genetic evidence that DUH1 is the adapter protein that recognizes the Hac1 u p tail and targets the covalently attached protein for degradation.

    Effects of DUH1 on Hac1p abundance, synthesis, and turnover

    Having established that DUH1 is required for degron-dependent silencing of two different reporter genes, we returned to HAC1 itself. To detect Hac1p we introduced a 3xHA tag at the extreme N terminus of the endogenous HAC1 allele (Figure 5A), which did not interfere with its post-transcriptional silencing in the absence of the UPR (Figure 5B and C). To determine which Hac1p isoforms are regulated by DUH1, we constructed a set of HA-tagged strains that constitutively produced either Hac1 i p containing the 18-amino-acid exon 2–encoded tail (construct 3), Hac1 u p containing the 10-amino-acid intron-encoded tail (construct 5), or Hac1 Δtail p containing no tail at all (constructs 4 and 6). All of the mRNAs encoding HA-tagged Hac1p variants were expressed at similar levels in the presence versus the absence of DUH1 (Figure 5B). Strikingly, at the protein level only the abundance of Hac1 u p was affected by disruption of DUH1, increasing by

    five fold in the knock-out strain (Figure 5C). The increased abundance of Hac1 u p in the absence of DUH1 was not due to increased translation, as evidenced by deletion of DUH1 having no impact on ribosome density on any of the HA-tagged reporter mRNAs (Figure 5D). Instead, our results suggest that Duh1p specifically affects the turnover of Hac1 u p due to its 10-amino-acid tail, as was suggested by the results of our reporter experiments.

    Effects of DUH1 on expression and stability of Hac1p.

    (A) Design of 3xHA-tagged HAC1 mRNA variants. Constructs are depicted as in Figure 2B, with the location of the N-terminal 3xHA tag indicated. (B) RNA abundance measurements for HAC1 mRNA variants. Total RNA was extracted from strains expressing the indicated mRNAs in either a DUH1 or duh1∆ background. qRT-PCR was used to measure the abundances of HAC1 variants relative to ACT1 mRNA, with all data normalized to the abundance of construct 1 in strain BY4741. Shown are the mean ± SD (n = 2). (C) Effect of DUH1 disruption on protein abundances. Strains expressing the indicated mRNAs depicted in (A) in either a DUH1 or duh1∆ background were grown to mid-log phase. Extracts were prepared and immunoblotted for 3xHA-Hac1p and actin loading control. (D) Polysome analysis of 3xHA-tagged HAC1 mRNA variants. Extracts were prepared in heparin-containing lysis buffer from strains expressing the mRNAs indicated in (A) in either a DUH1 or duh1∆ background. Polysome analysis was performed as in Figure 1A. (E) Analysis of protein degradation kinetics. Strains expressing construct 5 (depicted in A) in either a DUH1 or duh1∆ background were grown to mid-log phase before being treated with cycloheximide (CHX) to halt translation. At the indicated time points, aliquots of cells were quenched in dry-ice-cold methanol and harvested by centrifugation. Protein extraction and immunoblotting were performed as in (C), except that a high-sensitivity antibody was used to detect 3xHA-Hac1 u p. Shown are the mean ± SD (n = 3), expressed as a fraction of protein detected at t = 0.

    Because we were able to detect Hac1 u p by immunoblotting (using a high-sensitivity antibody) even when DUH1 was intact (unlike the corresponding GFP reporter), we could use cycloheximide (CHX) shut-off experiments to directly assay the impact of DUH1 on the turnover of Hac1 u p. In both the presence and absence of DUH1, Hac1 u p was degraded so rapidly that we could not accurately measure its half life even using a rapid harvesting procedure, due to the

    2 minutes required for CHX to accumulate in cells and halt translation (Gerashchenko and Gladyshev, 2014). However, the protein-degradation kinetics allowed us to calculate an upper bound for the half life of Hac1 u p, which was 50 seconds when DUH1 was present (Figure 5E). Deletion of DUH1 stabilized Hac1 u p and increased its half-life upper bound to 2 minutes. These results demonstrate that DUH1 is required for the extremely short half life of Hac1 u p that normally limits its accumulation. The true half-life difference upon DUH1 deletion is likely to be greater than the

    2-fold difference in upper bounds that we measured, based on the

    5-fold difference in steady-state protein levels that could not be accounted for by differences in either mRNA abundance or ribosome density (Figure 5B and D).

    Synergy between long-range base pairing and Duh1p-dependent degradation

    It was previously observed that disrupting the base-pairing interaction between the 5′-UTR and intron of HAC1 mRNA was sufficient to allow accumulation of Hac1 u p, which led to a model in which base-pairing alone was responsible for the post-transcriptional silencing phenomenon (Rüegsegger et al., 2001). Our results using constructs in which the base-pairing region was deleted (Figure 5) suggested that the previously observed accumulation of Hac1 u p was unknowingly being buffered by Duh1p-dependent degradation. To directly address this possibility, we generated HA-tagged constructs in which the base-pairing region was disrupted by mutations in either the 5′-UTR or intron or was reconstituted by the compensatory mutations (Figure 6A) and determined the effect of DUH1 deletion on steady-state protein levels. Because the 5′ and 3′ splice sites remained intact in these constructs, we introduced them into an ire1Δ background to eliminate any potential confounding effects of background splicing (Figure 1—figure supplement 1B). As previously observed, mutating the base-pairing region in the presence of DUH1 resulted in detectable levels of Hac1 u p (Figure 6C). However, the accumulation of Hac1 u p was greatly stimulated by deletion of DUH1, which was not explained by a corresponding increase in mRNA abundance (Figure 6B and C). Thus, although disrupting the base pairing produces detectable amounts of Hac1 u p as previously reported (Rüegsegger et al., 2001), Duh1p-dependent degradation restricts the steady-state level of the protein.

    Relationship between base pairing– and degron-dependent repression.

    (A) Design of 3xHA-tagged HAC1 mRNA variants. Constructs are depicted as in Figure 2B. Colored stars indicate mutations to the base-pairing region, with specific nucleotide changes shown in Figure 2B (red and blue stars) or below (green and black stars) in red. (B) RNA abundance measurements for HAC1 mRNA variants in the indicated strain backgrounds, analyzed as in Figure 5B. (CD) Effect of DUH1 disruption on protein abundances. ire1∆ strains expressing the indicated mRNAs depicted in (A) in either a DUH1 or duh1∆ background were analyzed as in Figure 5C, except that a high-sensitivity antibody was used to detect 3xHA-Hac1 u p.

    Remarkably, even a single nucleotide change in the center of the base-pairing region (Sathe et al., 2015) was sufficient for some accumulation of Hac1 u p, which again was enhanced by deletion of DUH1 (Figure 6D). This result suggests that the 5′-UTR–intron base-pairing interaction is only marginally stable, which may be required for efficient dissociation of the intron after splicing (see Discussion).

    Functional consequences of incomplete silencing

    Collectively, we have shown that a pair of silencing mechanisms, one translational and the other post-translational, prevents spurious production of Hac1 u p in the absence of the UPR. The existence of such a fail-safe silencing mechanism implies that ectopic production of Hac1 u p has physiological consequences that negatively impact cellular fitness. However, a previous study suggested that Hac1 u p is itself not an active transcription factor because it lacks the activating 18-amino-acid tail found in Hac1 i p (Mori et al., 2000), raising the question as to why Hac1 u p accumulation would need to be tightly regulated. We hypothesized that Hac1 u p was in fact an active transcription factor but that in the previous study its accumulation had been prevented due to the experiments being performed in a DUH1 background. To test this hypothesis, we evaluated the ability of strains that constitutively produced either Hac1 i p, Hac1 u p, or Hac1 Δtail p to grow under conditions of chronic ER stress induced by the drug tunicamycin. Strains expressing Hac1 i p or Hac1 Δtail p grew on tunicamycin-containing medium regardless of whether DUH1 (or IRE1) was present (Figure 7A), consistent with both proteins being active transcription factors that are not targeted by Duh1p. In contrast, strains expressing Hac1 u p only grew robustly on tunicamycin-containing medium when DUH1 was knocked out (but independently of IRE1). These results confirm our hypothesis that Duh1p-dependent degradation normally masks the activity of Hac1 u p. Our findings also explain how HAC1 was able to be initially identified as a high-copy activator of the UPR in an Δire1 strain, since the UPR activity detected in this strain had to have resulted from Hac1 u p produced from unspliced HAC1 mRNA (Chapman and Walter, 1997 Cox and Walter, 1996).

    Requirement for DUH1 to suppress Ire1p-independent activation of the UPR.

    (A) Analysis of Hac1p activity in the UPR. Strains expressing the indicated HAC1 mRNA variants, with (+) or without () IRE1 and/or DUH1 present, were grown to saturation. 10-fold dilution series were plated on YPD without (–Tm) or with (+Tm) 400 ng/ml tunicamycin to induce ER stress. (B) Impact of DUH1 on detection of Hac1 u p. Strains expressing the indicated mRNAs, with (+) or without () IRE1 and/or DUH1 present, were analyzed as in Figure 6C. Black arrow indicates the position of Hac1 u p, which migrates more slowly than Hac1 i p. Construct 1, which lacks a 3xHA tag, was used as a negative control for anti-HA immunoblotting. (C) Effect of Duh1p-dependent degradation on the UPR. Strains expressing wild-type HAC1 with an N-terminal 3xHA tag, with (+) or without () IRE1 and/or DUH1 present, were grown to saturation. 10-fold dilution series were plated on YPD without tunimacyin (–Tm) or containing the indicated concentration of tunicamycin (+Tm). Plates were imaged at days 2 (top), 3 (middle), and 6 (bottom).

    Because Hac1 u p has UPR-inducing activity (Figure 7A), we reasoned that the fail-safe mechanism we discovered is required to prevent leaky production of Hac1 u p that would otherwise cause Ire1p-independent activation of the UPR. However, we initially failed to detect Hac1 u p produced from unspliced HAC1 mRNA even when DUH1 was disrupted (Figure 5C). On the other hand, our results from the analogous GFP and HIS3 reporter gene studies demonstrated that translational repression mediated by 5′-UTR–intron base pairing was incomplete and allowed a low level of protein synthesis that was normally 'cleaned up' by Duh1p-dependent degradation (Figure 4D and E). This led us to hypothesize that Hac1 u p itself was also being occasionally produced from HAC1 u mRNA and rapidly degraded in a Duh1p-dependent manner, but that the short half life of Hac1 u p (Figure 5E)—relative to both GFP (Natarajan et al., 1998) and His3p (Belle et al., 2006)—further reduced the steady-state abundance of Hac1 u p to an extremely low level that was initially undetectable.

    To enhance detection of HA-tagged Hac1 u p, we made two modifications: We used a more sensitive anti-HA antibody for immunoblotting and we knocked out IRE1 in our strains, which eliminates the background Ire1p-dependent splicing of HAC1 mRNA (Figure 1—figure supplement 1B) and concomitant production of Hac1 i p that can otherwise dominate the signal on immunoblots. With these modifications we could now detect Hac1 u p being produced from unspliced HAC1 mRNA, but only when the protein was stabilized by deletion of DUH1 (Figure 7B) and at levels far below even those of Hac1 u p being constitutively produced in the presence of DUH1 (Figure 7—figure supplement 1). Despite the relatively low level of leaky translation product we detected, these results provide molecular evidence that Hac1 u p is being continuously produced from HAC1 u mRNA but rapidly degraded due to its C-terminal degron.

    Does the leaky production of Hac1 u p unmasked by deletion of DUH1 have any functional consequences? The ability of constitutively produced Hac1 u p to promote survival under UPR-inducing conditions (Figure 7A) suggested that small amounts of Hac1 u p might also induce the UPR to some extent. To test this possibility, we examined how strains expressing HA-tagged but otherwise unmodified HAC1 mRNA grew on media containing different concentrations of tunicamycin. At the highest tunicamycin concentration only strains expressing IRE1 could grow regardless of whether DUH1 was also present (Figure 7C), suggesting that growth was dependent on the abundant Hac1 i p produced from HAC1 i mRNA. In contrast, at lower concentrations of tunicamycin the IRE1 deletion strain showed some growth, but this was reproducibly enhanced by simultaneous deletion of DUH1. These results indicate that the low level of Hac1 u p detectable in strains lacking DUH1 (Figure 7B) is sufficient to activate the UPR enough to facilitate cell survival under stress, even in the absence of Ire1p. Thus, degron-dependent degradation of Hac1 u p mediated by Duh1p is normally required to prevent ectopic Ire1p-independent activation of the UPR.


    Mechanisms of Antibacterial Drugs

    Learning Objective

    Describe the mechanisms of action associated with drugs that inhibit cell wall biosynthesis, protein synthesis, membrane function, nucleic acid synthesis, and metabolic pathways

    An important quality for an antimicrobial drug is selective toxicity , meaning that it selectively kills or inhibits the growth of microbial targets while causing minimal or no harm to the host. Most antimicrobial drugs currently in clinical use are antibacterial because the prokaryotic cell provides a greater variety of unique targets for selective toxicity, in comparison to fungi, parasites, and viruses. Each class of antibacterial drugs has a unique mode of action (the way in which a drug affects microbes at the cellular level), and these are summarized in Figure 14.9 and Table 14.1 .

    Vaz, L.E., et al. “Prevalence of Parental Misconceptions About Antibiotic Use.” Pediatrics 136 no.2 (August 2015). DOI: 10.1542/ peds.2015-0883.

    Figure 14.9 There are several classes of antibacterial compounds that are typically classified based on their bacterial target.

    Common Antibacterial Drugs by Mode of Action

    Inhibitors of Cell Wall Biosynthesis

    Several different classes of antibacterials block steps in the biosynthesis of peptidoglycan, making cells more susceptible to osmotic lysis ( Table 14.2 ). Therefore, antibacterials that target cell wall biosynthesis are bactericidal in their action. Because human cells do not make peptidoglycan, this mode of action is an excellent example of selective toxicity.

    Penicillin, the first antibiotic discovered, is one of several antibacterials within a class called β-lactams . This group of compounds includes the penicillins, cephalosporins, monobactams, and carbapenems, and is characterized by the presence of a β-lactam ring found within the central structure of the drug molecule ( Figure 14.10 ). The β- lactam antibacterials block the crosslinking of peptide chains during the biosynthesis of new peptidoglycan in the bacterial cell wall. They are able to block this process because the β-lactam structure is similar to the structure of the peptidoglycan subunit component that is recognized by the crosslinking transpeptidase enzyme, also known as a penicillin-binding protein (PBP). Although the β-lactam ring must remain unchanged for these drugs to retain their antibacterial activity, strategic chemical changes to the R groups have allowed for development of a wide variety of semisynthetic β-lactam drugs with increased potency, expanded spectrum of activity, and longer half-lives for better dosing, among other characteristics.

    Penicillin G and penicillin V are natural antibiotics from fungi and are primarily active against gram-positive bacterial pathogens, and a few gram-negative bacterial pathogens such as Pasteurella multocida . Figure 14.10 summarizes the semisynthetic development of some of the penicillins. Adding an amino group (-NH 2 ) to penicillin G created the aminopenicillins (i.e., ampicillin and amoxicillin) that have increased spectrum of activity against more gram- negative pathogens. Furthermore, the addition of a hydroxyl group (-OH) to amoxicillin increased acid stability, which allows for improved oral absorption. Methicillin is a semisynthetic penicillin that was developed to address the spread of enzymes (penicillinases) that were inactivating the other penicillins. Changing the R group of penicillin G to the more bulky dimethoxyphenyl group provided protection of the β-lactam ring from enzymatic destruction by penicillinases, giving us the first penicillinase-resistant penicillin.

    Similar to the penicillins, cephalosporins contain a β-lactam ring ( Figure 14.10 ) and block the transpeptidase activity of penicillin-binding proteins. However, the β-lactam ring of cephalosporins is fused to a six-member ring, rather than the five-member ring found in penicillins. This chemical difference provides cephalosporins with an increased resistance to enzymatic inactivation by β-lactamases . The drug cephalosporin C was originally isolated from the fungus Cephalosporium acremonium in the 1950s and has a similar spectrum of activity to that of penicillin against gram-positive bacteria but is active against more gram-negative bacteria than penicillin. Another important structural difference is that cephalosporin C possesses two R groups, compared with just one R group for penicillin, and this provides for greater diversity in chemical alterations and development of semisynthetic cephalosporins. The family of semisynthetic cephalosporins is much larger than the penicillins, and these drugs have been classified into generations based primarily on their spectrum of activity, increasing in spectrum from the narrow-spectrum, first-generation cephalosporins to the broad-spectrum, fourth-generation cephalosporins. A new fifth-generation cephalosporin has been developed that is active against methicillin-resistant Staphylococcus aureus (MRSA).

    The carbapenems and monobactams also have a β-lactam ring as part of their core structure, and they inhibit the transpeptidase activity of penicillin-binding proteins. The only monobactam used clinically is aztreonam. It is a narrow-spectrum antibacterial with activity only against gram-negative bacteria. In contrast, the carbapenem family includes a variety of semisynthetic drugs (imipenem, meropenem, and doripenem) that provide very broad-spectrum activity against gram-positive and gram-negative bacterial pathogens.

    The drug vancomycin , a member of a class of compounds called the glycopeptides , was discovered in the 1950s as a natural antibiotic from the actinomycete Amycolatopsis orientalis . Similar to the β-lactams, vancomycin inhibits cell wall biosynthesis and is bactericidal. However, in contrast to the β-lactams, the structure of vancomycin is not similar to that of cell-wall peptidoglycan subunits and does not directly inactivate penicillin-binding proteins. Rather, vancomycin is a very large, complex molecule that binds to the end of the peptide chain of cell wall precursors, creating a structural blockage that prevents the cell wall subunits from being incorporated into the growing N-acetylglucosamine and N-acetylmuramic acid (NAM-NAG) backbone of the peptidoglycan structure (transglycosylation). Vancomycin also structurally blocks transpeptidation. Vancomycin is bactericidal against gram-

    positive bacterial pathogens, but it is not active against gram-negative bacteria because of its inability to penetrate the protective outer membrane.

    The drug bacitracin consists of a group of structurally similar peptide antibiotics originally isolated from Bacillus subtilis . Bacitracin blocks the activity of a specific cell-membrane molecule that is responsible for the movement of peptidoglycan precursors from the cytoplasm to the exterior of the cell, ultimately preventing their incorporation into the cell wall. Bacitracin is effective against a wide range of bacteria, including gram-positive organisms found on the skin, such as Staphylococcus and Streptococcus . Although it may be administered orally or intramuscularly in some circumstances, bacitracin has been shown to be nephrotoxic (damaging to the kidneys). Therefore, it is more commonly combined with neomycin and polymyxin in topical ointments such as Neosporin.

    Figure 14.10 Penicillins, cephalosporins, monobactams, and carbapenems all contain a β-lactam ring, the site of attack by inactivating β-lactamase enzymes. Although they all share the same nucleus, various penicillins differ from each other in the structure of their R groups. Chemical changes to the R groups provided increased spectrum of activity, acid stability, and resistance to β-lactamase degradation.

    Drugs that Inhibit Bacterial Cell Wall Synthesis

    Mechanism of Action

    Natural or Semisynthetic

    Spectrum of Activity

    Narrow-spectrum against gram-positive and a few gram-negative bacteria

    Drugs that Inhibit Bacterial Cell Wall Synthesis

    Mechanism of Action

    Natural or Semisynthetic

    Spectrum of Activity

    Narrow-spectrum against gram-positive bacteria but with increased gram- negative spectrum

    Narrow-spectrum against gram-positive bacteria only, including strains producing penicillinase

    Narrow-spectrum similar to penicillin but with increased gram-negative spectrum

    First- generation cephalosporins

    Narrow-spectrum similar to cephalosporin C

    Second- generation cephalosporins

    Narrow-spectrum but with increased gram-negative spectrum compared with first generation

    Third- and fourth- generation cephalosporins

    Broad-spectrum against gram-positive and gram- negative bacteria, including some β- lactamase producers

    Fifth- generation cephalosporins

    Broad-spectrum against gram-positive and gram- negative bacteria, including MRSA

    Narrow-spectrum against gram-negative bacteria, including some β- lactamase producers

    Broadest spectrum of the β-lactams against gram- positive and gram- negative bacteria, including many β- lactamase producers

    Narrow spectrum against gram-positive bacteria only, including multidrug- resistant strains

    Drugs that Inhibit Bacterial Cell Wall Synthesis

    Mechanism of Action

    Natural or Semisynthetic

    Spectrum of Activity

    Describe the mode of action of β-lactams.

    Inhibitors of Protein Biosynthesis

    The cytoplasmic ribosomes found in animal cells (80S) are structurally distinct from those found in bacterial cells (70S), making protein biosynthesis a good selective target for antibacterial drugs. Several types of protein biosynthesis inhibitors are discussed in this section and are summarized in Figure 14.11 .

    Protein Synthesis Inhibitors That Bind the 30S Subunit

    Aminoglycosides are large, highly polar antibacterial drugs that bind to the 30S subunit of bacterial ribosomes, impairing the proofreading ability of the ribosomal complex. This impairment causes mismatches between codons and anticodons, resulting in the production of proteins with incorrect amino acids and shortened proteins that insert into the cytoplasmic membrane. Disruption of the cytoplasmic membrane by the faulty proteins kills the bacterial cells. The aminoglycosides , which include drugs such as streptomycin, gentamicin, neomycin, and kanamycin, are potent broad-spectrum antibacterials. However, aminoglycosides have been shown to be nephrotoxic (damaging to kidney), neurotoxic (damaging to the nervous system), and ototoxic (damaging to the ear).

    Another class of antibacterial compounds that bind to the 30S subunit is the tetracyclines . In contrast to aminoglycosides, these drugs are bacteriostatic and inhibit protein synthesis by blocking the association of tRNAs with the ribosome during translation. Naturally occurring tetracyclines produced by various strains of Streptomyces were first discovered in the 1940s, and several semisynthetic tetracyclines, including doxycycline and tigecycline have also been produced. Although the tetracyclines are broad spectrum in their coverage of bacterial pathogens, side effects that can limit their use include phototoxicity, permanent discoloration of developing teeth, and liver toxicity with high doses or in patients with kidney impairment.

    Protein Synthesis Inhibitors That Bind the 50S Subunit

    There are several classes of antibacterial drugs that work through binding to the 50S subunit of bacterial ribosomes. The macrolide antibacterial drugs have a large, complex ring structure and are part of a larger class of naturally produced secondary metabolites called polyketides, complex compounds produced in a stepwise fashion through the repeated addition of two-carbon units by a mechanism similar to that used for fatty acid synthesis. Macrolides are broad-spectrum, bacteriostatic drugs that block elongation of proteins by inhibiting peptide bond formation between specific combinations of amino acids. The first macrolide was erythromycin . It was isolated in 1952 from Streptomyces erythreus and prevents translocation. Semisynthetic macrolides include azithromycin and telithromycin. Compared with erythromycin, azithromycin has a broader spectrum of activity, fewer side effects, and a significantly longer half-life (1.5 hours for erythromycin versus 68 hours for azithromycin) that allows for once-daily dosing and a short 3-day course of therapy (i.e., Zpac formulation) for most infections. Telithromycin is the first semisynthetic

    within the class known as ketolides. Although telithromycin shows increased potency and activity against macrolide- resistant pathogens, the US Food and Drug Administration (FDA) has limited its use to treatment of community- acquired pneumonia and requires the strongest “black box warning” label for the drug because of serious hepatotoxicity.

    The lincosamides include the naturally produced lincomycin and semisynthetic clindamycin . Although structurally distinct from macrolides, lincosamides are similar in their mode of action to the macrolides through binding to the 50S ribosomal subunit and preventing peptide bond formation. Lincosamides are particularly active against streptococcal and staphylococcal infections.

    The drug chloramphenicol represents yet another structurally distinct class of antibacterials that also bind to the 50S ribosome, inhibiting peptide bond formation. Chloramphenicol, produced by Streptomyces venezuelae , was discovered in 1947 in 1949, it became the first broad-spectrum antibiotic that was approved by the FDA. Although it is a natural antibiotic, it is also easily synthesized and was the first antibacterial drug synthetically mass produced. As a result of its mass production, broad-spectrum coverage, and ability to penetrate into tissues efficiently, chloramphenicol was historically used to treat a wide range of infections, from meningitis to typhoid fever to conjunctivitis. Unfortunately, serious side effects, such as lethal gray baby syndrome, and suppression of bone marrow production, have limited its clinical role. Chloramphenicol also causes anemia in two different ways. One mechanism involves the targeting of mitochondrial ribosomes within hematopoietic stem cells, causing a reversible, dose-dependent suppression of blood cell production. Once chloramphenicol dosing is discontinued, blood cell production returns to normal. This mechanism highlights the similarity between 70S ribosomes of bacteria and the 70S ribosomes within our mitochondria. The second mechanism of anemia is idiosyncratic (i.e., the mechanism is not understood), and involves an irreversible lethal loss of blood cell production known as aplastic anemia. This mechanism of aplastic anemia is not dose dependent and can develop after therapy has stopped. Because of toxicity concerns, chloramphenicol usage in humans is now rare in the United States and is limited to severe infections unable to be treated by less toxic antibiotics. Because its side effects are much less severe in animals, it is used in veterinary medicine.

    The oxazolidinones , including linezolid, are a new broad-spectrum class of synthetic protein synthesis inhibitors that bind to the 50S ribosomal subunit of both gram-positive and gram-negative bacteria. However, their mechanism of action seems somewhat different from that of the other 50S subunit-binding protein synthesis inhibitors already discussed. Instead, they seem to interfere with formation of the initiation complex (association of the 50S subunit, 30S subunit, and other factors) for translation, and they prevent translocation of the growing protein from the ribosomal A site to the P site. Table 14.3 summarizes the protein synthesis inhibitors.

    Figure 14.11 The major classes of protein synthesis inhibitors target the 30S or 50S subunits of cytoplasmic ribosomes.

    Drugs That Inhibit Bacterial Protein Synthesis

    Mechanism of Action

    Bacteriostatic or Bactericidal

    Spectrum of Activity

    Causes mismatches between codons and anticodons, leading to faulty proteins that insert into and disrupt cytoplasmic membrane Aminoglycosides Streptomycin, gentamicin, neomycin, kanamycin Blocks peptide bond formation between amino acids Macrolides Erythromycin, azithromycin, telithromycin

    Compare and contrast the different types of protein synthesis inhibitors.

    Inhibitors of Membrane Function

    A small group of antibacterials target the bacterial membrane as their mode of action ( Table 14.4 ). The polymyxins are natural polypeptide antibiotics that were first discovered in 1947 as products of Bacillus polymyxa only polymyxin B and polymyxin E ( colistin ) have been used clinically. They are lipophilic with detergent-like properties and interact with the lipopolysaccharide component of the outer membrane of gram-negative bacteria, ultimately disrupting both their outer and inner membranes and killing the bacterial cells. Unfortunately, the membrane-targeting mechanism is not a selective toxicity, and these drugs also target and damage the membrane of cells in the kidney and nervous system when administered systemically. Because of these serious side effects and their poor absorption from the digestive tract, polymyxin B is used in over-the-counter topical antibiotic ointments (e.g., Neosporin), and oral colistin was historically used only for bowel decontamination to prevent infections originating from bowel microbes in immunocompromised patients or for those undergoing certain abdominal surgeries. However, the emergence and spread of multidrug-resistant pathogens has led to increased use of intravenous colistin in hospitals, often as a drug of last resort to treat serious infections. The antibacterial daptomycin is a cyclic lipopeptide produced by Streptomyces roseosporus that seems to work like the polymyxins, inserting in the bacterial cell membrane and disrupting it. However, in contrast to polymyxin B and colistin, which target only gram-negative bacteria, daptomycin specifically targets gram-positive bacteria. It is typically administered intravenously and seems to be well tolerated, showing reversible toxicity in skeletal muscles.

    Drugs That Inhibit Bacterial Membrane Function

    Mechanism of Action

    Spectrum of Activity

    Intravenous dosing to treat serious systemic

    Drugs That Inhibit Bacterial Membrane Function

    Mechanism of Action

    Spectrum of Activity

    How do polymyxins inhibit membrane function?

    Inhibitors of Nucleic Acid Synthesis

    Some antibacterial drugs work by inhibiting nucleic acid synthesis ( Table 14.5 ). For example, metronidazole is a semisynthetic member of the nitroimidazole family that is also an antiprotozoan. It interferes with DNA replication in target cells. The drug rifampin is a semisynthetic member of the rifamycin family and functions by blocking RNA polymerase activity in bacteria. The RNA polymerase enzymes in bacteria are structurally different from those in eukaryotes, providing for selective toxicity against bacterial cells. It is used for the treatment of a variety of infections, but its primary use, often in a cocktail with other antibacterial drugs, is against mycobacteria that cause tuberculosis. Despite the selectivity of its mechanism, rifampin can induce liver enzymes to increase metabolism of other drugs being administered (antagonism), leading to hepatotoxicity (liver toxicity) and negatively influencing the bioavailability and therapeutic effect of the companion drugs.

    One member of the quinolone family, a group of synthetic antimicrobials, is nalidixic acid . It was discovered in 1962 as a byproduct during the synthesis of chloroquine, an antimalarial drug. Nalidixic acid selectively inhibits the activity of bacterial DNA gyrase, blocking DNA replication. Chemical modifications to the original quinolone backbone have resulted in the production of fluoroquinolones , like ciprofloxacin and levofloxacin, which also inhibit the activity of DNA gyrase. Ciprofloxacin and levofloxacin are effective against a broad spectrum of gram-positive or gram-negative bacteria, and are among the most commonly prescribed antibiotics used to treat a wide range of infections, including urinary tract infections, respiratory infections, abdominal infections, and skin infections. However, despite their selective toxicity against DNA gyrase, side effects associated with different fluoroquinolones include phototoxicity, neurotoxicity, cardiotoxicity, glucose metabolism dysfunction, and increased risk for tendon rupture.

    Drugs That Inhibit Bacterial Nucleic Acid Synthesis

    Mechanisms of Action

    Spectrum of activity

    Narrow spectrum with activity

    against gram-positive and limited

    numbers of gram-negative bacteria.

    positive and gram-negative bacteria

    Why do inhibitors of bacterial nucleic acid synthesis not target host cells?

    Inhibitors of Metabolic Pathways

    Some synthetic drugs control bacterial infections by functioning as antimetabolites , competitive inhibitors for bacterial metabolic enzymes ( Table 14.6 ). The sulfonamides ( sulfa drugs ) are the oldest synthetic antibacterial agents and are structural analogues of para -aminobenzoic acid (PABA), an early intermediate in folic acid synthesis ( Figure 14.12 ). By inhibiting the enzyme involved in the production of dihydrofolic acid, sulfonamides block bacterial biosynthesis of folic acid and, subsequently, pyrimidines and purines required for nucleic acid synthesis. This mechanism of action provides bacteriostatic inhibition of growth against a wide spectrum of gram-positive and gram-negative pathogens. Because humans obtain folic acid from food instead of synthesizing it intracellularly, sulfonamides are selectively toxic for bacteria. However, allergic reactions to sulfa drugs are common. The sulfones are structurally similar to sulfonamides but are not commonly used today except for the treatment of Hansen’s disease (leprosy).

    Trimethoprim is a synthetic antimicrobial compound that serves as an antimetabolite within the same folic acid synthesis pathway as sulfonamides. However, trimethoprim is a structural analogue of dihydrofolic acid and inhibits a later step in the metabolic pathway ( Figure 14.12 ). Trimethoprim is used in combination with the sulfa drug sulfamethoxazole to treat urinary tract infections, ear infections, and bronchitis. As discussed, the combination of trimethoprim and sulfamethoxazole is an example of antibacterial synergy. When used alone, each antimetabolite only decreases production of folic acid to a level where bacteriostatic inhibition of growth occurs. However, when used in combination, inhibition of both steps in the metabolic pathway decreases folic acid synthesis to a level that is lethal to the bacterial cell. Because of the importance of folic acid during fetal development, sulfa drugs and trimethoprim use should be carefully considered during early pregnancy.

    The drug isoniazid is an antimetabolite with specific toxicity for mycobacteria and has long been used in combination with rifampin or streptomycin in the treatment of tuberculosis. It is administered as a prodrug, requiring activation through the action of an intracellular bacterial peroxidase enzyme, forming isoniazid-nicotinamide adenine dinucleotide (NAD) and isoniazid-nicotinamide adenine dinucleotide phosphate (NADP), ultimately preventing the synthesis of mycolic acid, which is essential for mycobacterial cell walls. Possible side effects of isoniazid use include hepatotoxicity, neurotoxicity, and hematologic toxicity (anemia).

    Figure 14.12 Sulfonamides and trimethoprim are examples of antimetabolites that interfere in the bacterial synthesis of folic acid by blocking purine and pyrimidine biosynthesis, thus inhibiting bacterial growth.

    Antimetabolite Drugs

    Metabolic Pathway Target

    Mechanism of Action

    Spectrum of Activity

    Broad spectrum against gram-positive and gram- negative bacteria

    Broad spectrum against gram-positive and gram- negative bacteria

    Antimetabolite Drugs

    Metabolic Pathway Target

    Mechanism of Action

    Spectrum of Activity

    Narrow spectrum against Mycobacterium spp., including M. tuberculosis

    How do sulfonamides and trimethoprim selectively target bacteria?

    Inhibitor of ATP Synthase

    Bedaquiline, representing the synthetic antibacterial class of compounds called the diarylquinolones, uses a novel mode of action that specifically inhibits mycobacterial growth. Although the specific mechanism has yet to be elucidated, this compound appears to interfere with the function of ATP synthases, perhaps by interfering with the use of the hydrogen ion gradient for ATP synthesis by oxidative phosphorylation, leading to reduced ATP production. Due to its side effects, including hepatotoxicity and potentially lethal heart arrhythmia, its use is reserved for serious, otherwise untreatable cases of tuberculosis.


    CONCLUDING REMARKS

    EF-P was first discovered as an elongation factor that promoted peptide bond formation between poor aminoacyl acceptors. For a time, EF-P was misunderstood to be involved in the synthesis of the first peptide bond, but now we recognize it as a bona fide elongation factor, albeit a very unusual one in that it exhibits sequence specificity. Molecular-genetic and systems-level approaches show that the specificity requirement hangs on improving the translation rate through primary sequences encoding particularly poor peptidyl transfer substrates like consecutive prolines. Thus, after nearly 4 decades of study, our understanding of the activity of EF-P has come full circle and the role of EF-P in translation proposed by Glick, Chládek and Ganoza in 1979 appears to be accurate.

    While we understand a great deal more about EF-P today than when it was discovered, many aspects of EF-P biology remain mysterious. For example, EF-P promotes translation through many but not all XPPX motifs hinting at still further rules that govern its specificity. Moreover, not all proteins that experience XPPX-mediated stalling actually experience reduced protein copy number (Elgamal et al. 2014 Woolstenhulme et al. 2015). Exactly why some XPPX motifs are particularly difficult to translate isn't entirely clear, nor is the mechanism by which EF-P relieves this problem. The function of post-translational modification, once thought to be essential for EF-P activity, warrants reconsideration as unmodified EF-P and variants modified with a foreign chemical moiety have been recently reported as functional (Hummels et al. 2017 Volkwein et al. 2019). If the modification is part of the mechanism by which EF-P relieves translational pausing, how can so many diverse modifications all accommodate a conserved function? Alternatively, if the modification is regulatory, when and how is the modification added or altered to change EF-P function?

    Finally, and perhaps the biggest question, is: how does EF-P promote growth? The absence of EF-P results in reduced levels of a subset of proteins some of which are enzymes involved in essential metabolism. Enzymes are rather resistant to fluctuation in abundance, however, and it is not clear why a reduction in their level(s) would be growth limiting or even inhibitory. Moreover, recent work suggests that the requirement of EF-P, at least in E. coli, can be conditionally relieved simply by inducing slower growth at lower temperatures (Tollerson, Witzky and Ibba 2018). If slow growth and the corresponding reduction in translation rate is sufficient to abolish the need for EF-P, then why does EF-P appear to be essential in slow-growing organisms such as Mycobacterium (Sassetti, Boyd and Rubin 2003)? It is clear that EF-P is conserved in both form and function in all organisms across every domain of life and plays a specific but nonetheless important role in maintaining high rates of translation. Growth essentiality would be a strong selection to ensure such high conservation, but if EF-P is not strictly required for growth, why is it so highly conserved?


    Watch the video: Die Transkription - Proteinbiosynthese Teil 1 (August 2022).