Rapid detection of Escherichia coli in bloodstream infection via CRISPR-Cas9 engineered reporter phage T7::Nluc and microfluidic chip platform

Abstract

Rapid identification of pathogens responsible for bloodstream infection is critical for early intervention and effective treatment. Reporter phages, which are known for their exceptional sensitivity and specificity in pathogen detection, have garnered significant interest. In this study, we systematically evaluate phage genome editing strategies that combine homologous recombination with the CRISPR-Cas9 system. We investigate the impacts of homologous arm length, sgRNA activity, target site, and plasmid interactions on editing efficiency. Our results demonstrate that successful genome editing depends on both sufficient cleavage pressure and optimal homologous arm length, particularly when using low-activity sgRNAs. On the basis of these findings, we develop a highly efficient gene editing strategy TPMSR (triple-plasmid-mediated synchronous recombination) that overcomes the limitations of conventional methods that rely on high-activity sgRNA and restricted editing sites. Using the TPMSR strategy, we integrate the Nluc gene into phage T7, generating the reporter phage T7::Nluc, which is then incorporated into a microfluidic chip. Validation with 51 clinical isolates demonstrates outstanding sensitivity, specificity, and accuracy in detecting Escherichia coli in blood within 1.5 h at concentrations less than 30 CFU/mL. This study presents a robust strategy for phage genome engineering and develops a promising method for the rapid diagnosis of bloodstream infections caused by E. coli.

1. Introduction

Escherichia coli is a prevalent pathogen implicated in bloodstream infections (BSIs) [1], but traditional identification through blood culture and biochemical tests are time-consuming, typically requiring 24–48 h. Such delays are impractical in critical clinical situations. Although advanced techniques, including proteomics [2], high-throughput sequencing, and real-time PCR, have accelerated pathogen detection [3], these methods are costly and unable to distinguish viable organisms from nonviable organisms. Therefore, there is a critical need for a rapid, accurate and cost-effective method to identify pathogens, facilitate timely treatment, and minimize the inappropriate use of antibiotics.

Bacteriophages, the most diverse entities in the biosphere, offer a rich reservoir of host-specific tools, as nearly every bacterial species has a corresponding phage [4,5]. Exceptional exhibiting host specificity, ease of propagation, and simple structure, phages are ideal candidates for genetic engineering, particularly for constructing phage-based probes for pathogen detection [6]. Engineered phages expressing heterologous reporter genes (e.g., EGFP, Nluc, or ALP) can enable rapid identification of bacterial targets through protein expression [7–10]. However, traditional phage genome editing via homologous recombination is inefficient because of low recombination rates and the absence of selection markers, necessitating labor-intensive screening of recombinants [11]. Several strategies have been developed to increase phage editing efficiency, including the use of recombinases [12], whole-genome synthesis via yeast platforms [13], and Gibson assembly guided genome construction [14]. Despite these advancements, limitations such as technical complexity, instability, and the risk of introducing point mutations remain major obstacles to routine and reliable phage engineering.

The discovery of clustered regularly interspaced short palindromic repeats (CRISPR) has enabled the development of CRISPR-Cas9-based phage counterselection systems, which utilize sgRNA-directed, site-specific DNA cleavage [15]. This system relies on two key components: the sgRNA-Cas9 protein complex for DNA targeting and cleavage and the protospacer adjacent motif (PAM) sequence (NGG) for recognition [16,17]. By employing the CRISPR-Cas9 system, precise cleavage of the wild-type phage is achievable [18,19]. The introduction of silent mutations and deletions of PAM sites by homologous recombination enables recombinants to escape from CRISPR-Cas9 cleavage and facilitates the enrichment of recombinants [20–22]. However, this method is constrained by several factors, such as phage genome modifications (e.g., ghmC) [23,24], the location of cleavage sites [21,25], and homologous arm (HA) length [26,27], all of which can significantly influence editing efficiency [28]. To minimize functional disruption of phages, engineering in noncoding regions of genomes is often preferred. Nonetheless, identifying suitable sgRNAs in these regions poses a major challenge: while sgRNAs targeting coding regions are generally expected to exhibit higher activity in theory, empirical screening remains necessary.