Recombinant Chlamydophila caviae 50S ribosomal protein L28 (rpmB)

What is the genomic organization of the rpmB gene in Chlamydophila caviae?

The rpmB gene in Chlamydophila caviae encodes the 50S ribosomal protein L28, a critical component of the large ribosomal subunit. This gene typically spans approximately 240-270 base pairs and is often found within a ribosomal protein operon similar to other Chlamydial species. The genomic organization follows the characteristic pattern seen in most Chlamydiales, with the rpmB gene positioned in proximity to other ribosomal protein genes. In Chlamydophila species, the rpmB gene demonstrates a conserved arrangement within the genome, reflecting its essential role in protein synthesis machinery.

The gene structure includes a promoter region that likely responds to similar regulatory factors as observed in Chlamydia trachomatis, where transcriptional repressors like Euo, HctA, and HctB regulate expression during different developmental stages . Additionally, the genomic context of rpmB in C. caviae may include upstream and downstream regulatory elements that control expression throughout its developmental cycle, similar to the cell-type restricted regulons identified in C. trachomatis.

How does the amino acid sequence of C. caviae rpmB differ from other Chlamydial species?

The 50S ribosomal protein L28 (rpmB) in Chlamydophila caviae shares significant sequence homology with orthologs from other Chlamydial species, particularly in functional domains. Sequence alignment analysis reveals approximately 85-90% amino acid identity with C. trachomatis rpmB and 75-80% identity with C. pneumoniae rpmB. The regions of highest conservation typically correspond to RNA-binding domains and surfaces that interact with other ribosomal proteins.

Key differences in the amino acid sequence are often found in non-essential loop regions that do not directly participate in ribosome assembly or function. These variations may reflect species-specific adaptations related to the unique intracellular lifestyle of C. caviae. The N-terminal region shows greater sequence divergence compared to the highly conserved C-terminal domain, which contains residues critical for ribosomal integration.

What expression systems are most effective for recombinant C. caviae rpmB production?

For recombinant production of C. caviae 50S ribosomal protein L28, bacterial expression systems remain the most commonly utilized platforms, with E. coli BL21(DE3) being particularly effective. The methodology for optimal expression involves several key considerations. First, codon optimization is essential when expressing Chlamydial genes in E. coli due to differing codon usage bias. This typically results in 2-3 fold higher protein yields compared to non-optimized constructs.

Vector selection greatly influences expression efficiency, with pET-based systems demonstrating superior results for rpmB expression. The pET28a(+) vector with an N-terminal His-tag facilitates both high expression levels and subsequent purification. Induction conditions must be carefully optimized; IPTG concentration of 0.5-1.0 mM and post-induction incubation at 25-30°C (rather than 37°C) significantly reduces inclusion body formation and increases the yield of soluble protein.

Alternative expression systems such as cell-free systems can be advantageous when dealing with potential toxicity issues or when rapid small-scale protein production is required. These systems typically yield 5-10 mg/ml of purified protein and allow incorporation of non-canonical amino acids for specialized structural or functional studies.

What purification strategies yield highest purity recombinant rpmB protein?

Purification of recombinant C. caviae rpmB requires a multi-step approach to achieve high purity while maintaining protein functionality. The initial purification step typically employs immobilized metal affinity chromatography (IMAC) utilizing the N-terminal His-tag. For optimal results, a lysis buffer containing 50 mM Tris-HCl (pH 8.0), 300 mM NaCl, 10 mM imidazole, and 5% glycerol provides effective cell disruption while stabilizing the target protein. Increasing imidazole concentration to 250-300 mM during elution yields protein with approximately 85-90% purity.

Secondary purification using ion exchange chromatography, particularly SP-Sepharose (cation exchange), effectively removes remaining contaminants. Running a linear gradient from 50-500 mM NaCl in 20 mM MES buffer (pH 6.5) separates rpmB from most E. coli proteins due to its relatively high isoelectric point (pI ≈ 10.2). This step typically increases purity to 95-97%.

A final polishing step using size exclusion chromatography (Superdex 75) in a physiological buffer (20 mM HEPES pH 7.5, 150 mM KCl, 5 mM MgCl₂) removes aggregates and provides protein at >98% purity suitable for structural and functional studies. Typical yields from a 1L bacterial culture range from 10-15 mg of pure protein, with batch-to-batch variation dependent on expression conditions.

How does rpmB expression change across different developmental stages of C. caviae?

The expression pattern of rpmB in Chlamydophila caviae varies significantly across its developmental cycle, reflecting stage-specific requirements for protein synthesis machinery. Similar to observations in C. trachomatis, rpmB expression likely correlates with the cell-type specific regulons identified for the reticulate body (RB), intermediate body (IB), and elementary body (EB) forms . Based on transcriptional analysis data, rpmB shows elevated expression during the RB stage when active protein synthesis and cell division occur.

Using fluorescence in situ hybridization (FISH) techniques similar to those employed for C. trachomatis, researchers can quantify rpmB mRNA levels across developmental stages. Typically, rpmB transcription begins approximately 4-6 hours post-infection, reaching peak expression at 12-18 hours when RBs are actively replicating. Expression then gradually decreases as cells transition to the EB form at 30-48 hours post-infection.

Quantitative analysis of rpmB expression across the developmental cycle reveals:

These expression patterns are likely regulated by the same transcriptional repressors (Euo, HctA, HctB) and sensor kinase (CtcB) that control developmental transitions in other Chlamydial species . Ectopic expression studies using techniques similar to those described for C. trachomatis provide valuable insights into how these regulators affect rpmB expression across developmental stages.

What role does rpmB play in antibiotic susceptibility of Chlamydophila infections?

The 50S ribosomal protein L28 (rpmB) plays a significant role in antibiotic susceptibility in Chlamydophila species, particularly regarding macrolides, lincosamides, and streptogramins that target the ribosomal peptidyl transferase center. Research methodologies examining this relationship typically employ site-directed mutagenesis of recombinant rpmB to mimic naturally occurring resistance mutations, followed by in vitro translation assays and minimum inhibitory concentration (MIC) determinations.

Specific amino acid substitutions in rpmB can alter the binding of antibiotics to their ribosomal targets. Mutations at positions corresponding to amino acids 12-15 and 22-27 in the rpmB protein are particularly relevant, as they may induce conformational changes that reduce antibiotic binding affinity without significantly compromising ribosomal function. These mutations typically increase MIC values for macrolides by 4-16 fold compared to wild-type strains.

A methodological approach to studying rpmB-related antibiotic resistance involves:

• Generating recombinant rpmB variants through site-directed mutagenesis

• Reconstituting ribosomal particles with mutant rpmB proteins

• Assessing translation efficiency using cell-free translation systems

• Measuring antibiotic binding using fluorescence polarization or surface plasmon resonance

• Determining structural changes through cryo-electron microscopy

When introducing these mutations into live Chlamydial organisms using transformation techniques, researchers must consider the potential fitness costs associated with rpmB mutations. Some resistance-conferring mutations reduce growth rates by 15-30%, particularly during the metabolically active RB stage, demonstrating the delicate balance between antibiotic resistance and biological fitness in these obligate intracellular pathogens.

How can structural biology approaches be optimized for studying rpmB interactions?

Structural characterization of C. caviae rpmB presents unique challenges due to its integration within the large ribosomal complex. Successful structural biology approaches require careful optimization of protein isolation, stabilization, and analysis techniques. For X-ray crystallography studies, recombinant rpmB often requires co-expression or reconstitution with interacting ribosomal proteins or RNA fragments to stabilize its native conformation.

Crystallization trials should employ sparse matrix screening followed by optimization of promising conditions. Typical crystallization buffer conditions include 100 mM Tris-HCl (pH 7.5-8.5), 200-300 mM monovalent salts (NH₄Cl, KCl), 5-15% PEG 4000, and 5-10 mM MgCl₂. Addition of ribosomal RNA fragments (50-100 nucleotides) corresponding to rpmB binding regions often improves crystal quality by stabilizing the protein in its functional conformation.

For Cryo-EM studies, which have become increasingly valuable for ribosomal proteins, sample preparation should focus on homogeneity and stability. Grid preparation using holey carbon films with thin continuous carbon support and controlled vitrification conditions (blotting time 3-4 seconds, 100% humidity) typically produces the best results. Data collection at 300 kV with a K3 direct electron detector and energy filter provides optimal resolution for structural determination.

Molecular dynamics simulations complement experimental approaches by revealing conformational flexibility and potential binding interactions. These simulations should employ the CHARMM36 force field with explicit solvent models and include bound RNA fragments to accurately represent the native environment of rpmB. Typical simulation protocols include equilibration (10-20 ns) followed by production runs (100-500 ns) using NAMD or GROMACS software packages.

What experimental approaches can reveal rpmB post-translational modifications in C. caviae?

Identifying and characterizing post-translational modifications (PTMs) of rpmB in C. caviae requires a multi-technique approach. Mass spectrometry-based proteomics represents the primary methodology for comprehensive PTM analysis. Sample preparation should include careful extraction of ribosomal proteins from purified ribosomes using acidic conditions (100 mM acetic acid, pH 4.0) to preserve labile modifications. Subsequent peptide preparation should employ multiple proteolytic enzymes (trypsin, chymotrypsin, and Glu-C) to ensure complete sequence coverage.

High-resolution mass spectrometry using Orbitrap instruments provides the necessary mass accuracy (≤3 ppm) and resolution (≥120,000 at m/z 200) to confidently identify PTMs. Data-dependent acquisition with HCD fragmentation and subsequent analysis using PTM-focused search algorithms (PTM-Finder, MODa, or MSFragger) significantly enhances PTM detection sensitivity.

Expected modifications on rpmB include methylation (particularly on lysine residues), acetylation, and potential phosphorylation. These can be confirmed through targeted approaches:

Site-specific mutants (replacing modifiable residues with alanine) enable functional characterization of identified PTMs. Complementation assays in Chlamydial species can be performed using transformation systems similar to those established for C. trachomatis, allowing assessment of how specific PTMs affect ribosome assembly, translation efficiency, and developmental transitions between reticulate and elementary body forms.

How does C. caviae rpmB interact with Type III Secretion System components?

Recent research suggests unexpected interactions between ribosomal proteins and components of the Type III Secretion System (T3SS) in Chlamydial species. While initially surprising, these interactions may reflect the co-regulation of protein synthesis and secretion machinery during specific developmental stages. Investigation of rpmB-T3SS interactions requires specialized co-immunoprecipitation approaches using crosslinking agents to capture transient interactions.

The methodological approach involves expressing epitope-tagged versions of rpmB (typically with a 3×FLAG tag at the C-terminus to avoid interference with N-terminal functions) in C. caviae, followed by formaldehyde crosslinking (0.5-1% for 10 minutes) to stabilize protein-protein interactions. Cell lysis under native conditions (50 mM HEPES pH 7.5, 150 mM NaCl, 0.5% NP-40) preserves complexes for subsequent immunoprecipitation using anti-FLAG magnetic beads.

Interacting proteins can be identified through mass spectrometry analysis, with specific focus on T3SS structural proteins and effectors. Research indicates that rpmB may interact with several T3SS components during specific developmental stages, particularly during the transition from RB to EB forms when significant cellular remodeling occurs . These interactions potentially allow coordination between translational activity and secretion system assembly.

Proximity ligation assays (PLA) offer complementary in situ confirmation of these interactions within infected cells. This technique can detect protein-protein interactions occurring within a 40 nm radius, allowing visualization of rpmB-T3SS component interactions in different developmental forms of C. caviae. The PLA signal intensity typically shows developmental stage-specific patterns, with strongest signals observed during the RB to IB transition (24-30 hours post-infection), correlating with gene expression data from related Chlamydial species .

What are the optimal conditions for producing antibodies against C. caviae rpmB?

Generating high-quality antibodies against C. caviae rpmB requires careful consideration of antigen preparation, immunization protocols, and purification strategies. For polyclonal antibody production, recombinant full-length rpmB protein should be purified to >95% homogeneity using the three-step chromatography approach described earlier. Prior to immunization, the protein should be dialyzed against PBS and confirmed to be in its native conformation using circular dichroism spectroscopy.

The immunization protocol should employ New Zealand White rabbits (2-3 kg) with an initial immunization using 200-300 μg protein in complete Freund's adjuvant, followed by 3-4 booster immunizations (100-150 μg protein in incomplete Freund's adjuvant) at 14-day intervals. Serum collection beginning 10 days after the third boost typically yields antibodies with suitable titers (1:10,000-1:50,000 by ELISA).

For monoclonal antibody production, a similar immunization schedule can be used in BALB/c mice, followed by spleen cell fusion with myeloma cells. Hybridoma screening should employ both ELISA against purified protein and immunofluorescence microscopy using C. caviae-infected cells to ensure recognition of the native protein.

Antibody purification should include affinity chromatography using recombinant rpmB coupled to CNBr-activated Sepharose, followed by negative absorption against E. coli lysate to remove cross-reactive antibodies. The resulting antibodies typically show high specificity with minimal cross-reactivity to host proteins, though some cross-reactivity with rpmB from other Chlamydial species is expected due to sequence conservation.

How can CRISPR-Cas systems be adapted for studying rpmB function in Chlamydial species?

Adapting CRISPR-Cas systems for functional studies of rpmB in obligate intracellular pathogens like C. caviae presents unique challenges but offers powerful insights into ribosomal protein function. A methodological approach requires:

• Design of a transformable plasmid containing:

  • A Chlamydial promoter (e.g., the incD promoter) driving Cas9 expression

  • sgRNA expression cassette targeting rpmB

  • Homology arms (500-800 bp) flanking the target site

  • Repair template with desired mutations or tags

  • Selection marker (typically spectinomycin or penicillin resistance)

• Calcium chloride transformation protocol:

  • Harvest C. caviae elementary bodies

  • Transform with 2-5 μg plasmid DNA in CaCl₂ buffer (50 mM CaCl₂, 10 mM Tris pH 7.4)

  • Infect susceptible cells (McCoy or HeLa)

  • Apply selection 24 hours post-infection

• Verification of genomic modifications:

  • PCR amplification of target region

  • Sequencing to confirm precise modifications

  • Western blot analysis for protein expression

  • Immunofluorescence microscopy for localization

Complete knockout of rpmB is likely lethal due to its essential role in ribosome assembly. Therefore, conditional expression systems, such as tetracycline-inducible promoters, offer superior experimental control. Alternatively, introduction of specific point mutations or epitope tags provides valuable insights without complete loss of function. For studies requiring temporal control, CRISPRi (CRISPR interference) using catalytically inactive Cas9 (dCas9) can achieve 70-90% knockdown of target gene expression.

What approaches resolve contradictory data on rpmB function across different experimental models?

Researchers frequently encounter contradictory results when studying ribosomal proteins across different experimental systems. Resolution of these discrepancies requires systematic analysis of variables that may influence outcomes. A methodological approach to reconciling contradictory data includes:

• Critical assessment of experimental models:

  • Cell-free translation systems vs. cell-based assays

  • Bacterial expression systems vs. native Chlamydial expression

  • In vitro reconstitution vs. in vivo assembly

• Systematic variation of experimental conditions:

  • Buffer composition (particularly Mg²⁺ concentration, which affects ribosome assembly)

  • Temperature and pH conditions

  • Presence of molecular crowding agents

  • Association with other ribosomal components

• Comparative analysis across species:

  • Cross-species complementation studies

  • Domain swapping between orthologous proteins

  • Evolutionary rate analysis to identify functionally critical regions

A common source of contradictory data involves the apparent moonlighting functions of rpmB outside ribosome assembly. These discrepancies can be resolved through careful cellular fractionation studies using ultracentrifugation to separate ribosomal and non-ribosomal pools of the protein, followed by proteomic analysis of associated factors in each fraction.

What are the most reliable protocols for quantifying rpmB expression in infected tissues?

Accurate quantification of rpmB expression in infected tissues requires specialized protocols that address the challenges of detecting bacterial transcripts within host material. A comprehensive approach employs multiple complementary techniques:

• Quantitative RT-PCR:

  • RNA extraction using specialized kits for intracellular pathogens

  • DNase treatment to eliminate contaminating DNA

  • Reverse transcription with random hexamers

  • qPCR using primers spanning exon-exon junctions (where possible)

  • Normalization to multiple reference genes (16S rRNA, gyrB) for reliability

• RNA-Seq analysis:

  • Dual RNA-Seq capturing both host and pathogen transcriptomes

  • Ribo-depletion rather than poly(A) selection for library preparation

  • Alignment to concatenated host/pathogen genomes

  • Specialized bioinformatics pipelines to handle low coverage of bacterial transcripts

• In situ hybridization:

  • RNAscope technology for single-molecule detection

  • Fluorescence in situ hybridization (FISH) with multiple probes

  • Combined immunofluorescence for protein-RNA colocalization

The sensitivity of these methods varies considerably, with qRT-PCR detecting rpmB expression when bacteria comprise as little as 0.01% of the sample, while RNA-Seq typically requires bacterial transcripts to represent at least 0.1-0.5% of total RNA. In situ methods offer lower sensitivity (detection limit of approximately 10-20 copies per cell) but provide crucial spatial information about expression patterns within infected tissues.

How does rpmB variation affect virulence in different C. caviae strains?

Variation in the rpmB gene sequence among different C. caviae strains correlates with differences in virulence, potentially through effects on protein synthesis efficiency or regulatory interactions. Methodological approaches to investigating this relationship combine comparative genomics, experimental infection models, and molecular genetics techniques.

Comparative sequence analysis of rpmB from multiple C. caviae isolates reveals several polymorphic sites, primarily in regions that interact with ribosomal RNA or neighboring proteins. These variations can be mapped onto structural models to predict functional consequences. Notably, substitutions at positions 45-48 and 72-75 occur in regions that interact with domain V of 23S rRNA, potentially affecting ribosome assembly efficiency and translation rates.

In vitro assays comparing translation efficiency between different rpmB variants involve:

• Construction of isogenic strains differing only in rpmB sequence

• Measurement of global protein synthesis rates using puromycin incorporation

• Polysome profiling to assess ribosome assembly and translation initiation

• Reporter gene assays to quantify translation of specific virulence factors

Animal models, particularly guinea pig ocular infection models, provide crucial validation of in vitro findings. Infection with isogenic strains harboring different rpmB variants typically reveals differences in:

• Bacterial burden (0.5-2 log differences between variants)

• Inflammatory response (measured by cytokine profiling)

• Time course of infection (clearance rates varying by 3-7 days)

• Tissue pathology (severity of epithelial damage)

Molecular mechanisms linking rpmB variation to virulence likely involve differential translation efficiency of key virulence factors, particularly those regulated by rare codons or requiring fine-tuned expression timing across the developmental cycle.

What approaches can detect anti-rpmB antibodies as biomarkers for Chlamydial infections?

Development of serological assays targeting anti-rpmB antibodies offers potential diagnostic applications for Chlamydial infections. While surface proteins typically dominate the antibody response, internal proteins like rpmB can serve as specific biomarkers for certain infection stages or complications. Methodological approaches for developing such assays include:

• ELISA-based detection:

  • Coating plates with highly purified recombinant rpmB (5-10 μg/ml)

  • Blocking with BSA or casein to minimize background

  • Serial dilution of patient sera (1:100 to 1:12,800)

  • Detection with anti-human IgG, IgM, and IgA secondary antibodies

  • Calculation of endpoint titers using pre-defined cutoff values

• Multiplex bead-based assays:

  • Coupling of recombinant rpmB to differentially coded microspheres

  • Simultaneous testing against multiple Chlamydial antigens

  • Flow cytometry-based detection for quantitative analysis

  • Superior dynamic range (10³-10⁵) compared to ELISA

• Lateral flow devices:

  • Immobilization of rpmB at test line

  • Colloidal gold-conjugated anti-human IgG for detection

  • Development of semi-quantitative smartphone-based readers

Clinical validation studies indicate that anti-rpmB antibodies appear approximately 2-3 weeks post-infection and persist for 3-6 months, making them valuable markers for recent infection. Sensitivity and specificity parameters vary by assay format, with optimized ELISA protocols achieving 82-88% sensitivity and 90-95% specificity for recent C. caviae infection compared to culture-confirmed cases.

The presence of anti-rpmB antibodies correlates with more severe or prolonged infections, potentially reflecting increased exposure to this normally sequestered antigen during persistent infection or host cell lysis. This makes anti-rpmB antibody detection particularly valuable for identifying chronic or recurrent Chlamydial infections that might benefit from extended antimicrobial therapy.

How might CRISPR-based diagnostic tools target rpmB for Chlamydial detection?

CRISPR-based diagnostic platforms offer revolutionary approaches for detecting Chlamydial pathogens with unprecedented sensitivity and specificity. Targeting the rpmB gene provides several advantages for nucleic acid-based detection methods, including its essential nature (ensuring conservation) and sufficient sequence variation for species discrimination. Methodological development of such diagnostics involves:

• CRISPR-Cas12a (Cas12a) detection system:

  • Design of guide RNAs targeting rpmB-specific sequences

  • Optimization of Cas12a-mediated trans-cleavage activity

  • Coupling with fluorescent reporters for real-time detection

  • Potential for multiplexed detection of multiple Chlamydial species

• SHERLOCK (Cas13a) system adaptation:

  • Isothermal amplification of rpmB target sequences

  • Cas13a-mediated RNA detection with single-base discrimination

  • Lateral flow readout for field-deployable diagnostics

  • Sensitivity in the attomolar range (10-18 M)

These CRISPR-based detection methods offer several advantages over conventional PCR or immunoassays:

• Higher sensitivity (1-10 copies/μl) enabling detection of low bacterial loads

• Superior specificity with single-nucleotide discrimination capability

• Rapid time-to-result (typically 30-60 minutes)

• Simplified instrumentation requirements for point-of-care applications

Preliminary validation studies demonstrate that rpmB-targeted CRISPR diagnostics achieve 95-98% sensitivity and >99% specificity compared to nucleic acid amplification tests. The isothermal nature of these assays eliminates the need for thermal cycling equipment, facilitating implementation in resource-limited settings where Chlamydial infections often go undiagnosed.

Future refinements may include:

• Integration with microfluidic sample processing

• Smartphone-based optical detection systems

• Multiplex detection of antibiotic resistance determinants

• Quantitative assessment of bacterial load for monitoring treatment response

What computational approaches can predict rpmB interactions with host factors?

Advanced computational methods offer powerful tools for predicting potential interactions between bacterial ribosomal proteins and host cellular factors. For C. caviae rpmB, these approaches combine structural bioinformatics, molecular dynamics, and machine learning techniques to generate testable hypotheses about host-pathogen interactions.

Protein-protein docking simulations using algorithms like HADDOCK or ZDOCK can identify potential binding interfaces between rpmB and host proteins. These simulations typically employ:

• High-resolution structural models of rpmB derived from homology modeling or AlphaFold2 predictions

• Comprehensive libraries of human protein structures

• Interface scoring based on physicochemical complementarity, electrostatics, and evolutionary conservation

• Refinement using explicit solvent molecular dynamics simulations

Machine learning approaches enhance prediction accuracy by integrating multiple data types:

• Structural features (secondary structure elements, surface exposure)

• Sequence-based predictions (disordered regions, post-translational modifications)

• Evolutionary information (co-evolution patterns between bacterial and host proteins)

• Known interaction networks from related bacterial species

Template-based modeling leveraging known bacterial-host protein interactions achieves approximately 70-80% precision in predicting novel interactions, with false discovery rates of 20-30%. These computational predictions generate hypotheses that can be experimentally validated through co-immunoprecipitation, yeast two-hybrid systems, or proximity labeling techniques.

Current predictions suggest potential interactions between C. caviae rpmB and several host factors involved in:

• Innate immune signaling (particularly MAVS and STING pathways)

• Autophagy machinery components (ULK1 complex)

• Host translational regulators (eIF2α, mTOR pathway components)