Closed-Tube Colorimetric LAMP for Rapid Pathogen Detection in Suspected Neonatal Sepsis: Pre-Analytical Bottlenecks, Contamination Control, and Clinical Translation

1. Introduction

Neonatal sepsis remains a major cause of morbidity and mortality during the first 28 days of life, with a disproportionate burden among preterm and low-birth-weight infants and in healthcare settings with limited diagnostic resources [1,2]. Clinical recognition is difficult because the initial signs are often subtle, overlap with non-infectious neonatal conditions, and are interpreted using definitions that remain inconsistent across studies and clinical settings [3,4,5].

Blood culture remains the reference method for confirming bacterial or fungal sepsis [6]. Its diagnostic yield, however, depends strongly on the volume and quality of blood collected [7]. Inadequate inoculation can reduce sensitivity and complicate the interpretation of organisms that may represent contamination rather than true bloodstream infection [6]. Culture results are also rarely available early enough to guide the initial treatment decision [8,9]. Consequently, empirical antibiotics are frequently initiated before microbiological confirmation, exposing many infants without proven infection to unnecessary treatment [10,11].

Molecular assays can provide faster microbial detection [12], but their reported performance in neonatal sepsis remains heterogeneous [13]. A recent diagnostic-accuracy review estimated a summary sensitivity of 0.91 (95% confidence interval [CI], 0.85–0.95) and specificity of 0.88 (95% CI, 0.83–0.92), although the certainty of evidence was low and substantial variation persisted among platforms and study designs [13]. These findings support molecular assays as complementary tests rather than replacements for culture and antimicrobial susceptibility testing [3,13,14].

Loop-mediated isothermal amplification offers a practical alternative because it amplifies nucleic acids at a constant temperature and supports rapid visual detection without conventional thermal cycling [15,16]. Colorimetric readouts simplify result interpretation, while closed-tube detection limits post-amplification handling and reduces the opportunity for amplicon dispersal [17,18]. An extraction-free colorimetric LAMP assay recently detected Streptococcus agalactiae at 2 CFU/mL in spiked neonatal blood, demonstrating analytical promise without yet establishing diagnostic accuracy in prospectively enrolled neonates [19].

The clinical value of closed-tube colorimetric LAMP in neonatal sepsis depends not only on amplification chemistry but also on blood volume, recovery of intact pathogens, matrix-associated inhibition, contamination control, and reliable interpretation of negative and invalid results. Accordingly, this review evaluates the method as an integrated diagnostic pathway rather than as an amplification reaction in isolation. It brings together neonatal-blood preanalytics, assay and control design, pathogen-specific evidence, and clinical implementation while distinguishing analytical and simulated-matrix findings from direct clinical evidence. This framework is used to define the requirements for clinically interpretable testing alongside blood culture and to identify the validation steps needed before routine application in suspected neonatal sepsis.

2. Review Methodology and Evidence Framework

This structured narrative review used a transparent literature-search and evidence-mapping approach. Scopus, Web of Science Core Collection, and PubMed/MEDLINE were searched for English-language publications from January 2000 to 30 July Search terms combined “neonatal sepsis”, “neonatal bloodstream infection”, “loop-mediated isothermal amplification”, “LAMP”, “colorimetric”, “closed-tube”, “whole blood”, “sample preparation”, “inhibition”, “contamination”, and the names of selected neonatal-sepsis pathogens. Reference lists of relevant primary studies and reviews were also screened. Seminal pre-2000 studies were retained when they addressed foundational methods or neonatal blood-volume requirements.

Peer-reviewed studies were considered when they addressed LAMP-based pathogen detection, neonatal blood diagnostics, pre-analytical processing, amplification inhibition, contamination control, or clinical implementation. Animal, food, and environmental studies were included only when they provided an assay-design or contamination-control principle directly transferable to neonatal blood testing. Evidence was grouped as purified-template or isolate studies, spiked-matrix studies, clinical-specimen studies, prospective diagnostic-accuracy studies, or implementation studies. Sample matrix, blood volume, processing method, analytical sensitivity, assay time, comparator, diagnostic performance, and reported limitations were recorded where available.

The review was informed by SANRA principles, particularly transparent searching, appropriate referencing, and explicit differentiation between analytical and clinical evidence [20]. Because the available studies were heterogeneous in population, matrix, assay format, and comparator, findings were synthesized narratively rather than pooled quantitatively [13]. No formal risk-of-bias score was assigned.

3. Closed-Tube Colorimetric LAMP: Principles and Assay Design

Loop-mediated isothermal amplification (LAMP) amplifies a defined nucleic-acid target at a constant temperature using a strand-displacing DNA polymerase and a coordinated set of inner and outer primers [15,21]. The conventional reaction uses forward and backward inner primers, FIP and BIP, together with the outer primers F3 and B3 to recognize six regions of the target sequence and generate dumbbell-shaped intermediates that undergo continuous stem-loop amplification [21]. Loop primers provide additional initiation sites and can accelerate amplification without changing the basic reaction mechanism [22]. Most LAMP assays operate between 60 and 65 °C, although the optimum temperature and incubation period depend on the polymerase, target sequence, and primer set [15,23].

Assay design should begin with a target region that is conserved across the clinically relevant strains intended for detection and sufficiently distinct from closely related or commensal organisms. Whole-genome comparison can improve strain inclusivity and analytical exclusivity by screening candidate primer regions against both target and background genomes [24]. Candidate primer sets should then be evaluated experimentally using diverse target isolates, epidemiologically relevant non-target organisms, and replicated no-template controls because interactions among the multiple LAMP primers can generate nonspecific amplification [23,25,26]. Analytical sensitivity should be established with intact organisms carried through the intended neonatal-blood processing workflow rather than with purified nucleic acid alone [19,27]. A maximum incubation time should be fixed before clinical validation because late amplification may reflect primer-derived or non-template products [23,25]. For a multispecies panel using a common nonspecific color indicator, spatially separated sealed chambers are preferable to pooling multiple primer sets in one reaction, as compartmentalization preserves target attribution while maintaining a closed workflow [28,29].

Colorimetric LAMP converts chemical changes produced during amplification into a visible endpoint. Extensive DNA synthesis generates protons and pyrophosphate, alters the concentration of free magnesium ions, and produces large quantities of double-stranded DNA [16]. These changes can be detected indirectly using pH-sensitive indicators such as phenol red or metal-ion indicators such as hydroxy naphthol blue and calcein [16,30,31]. DNA-binding dyes report the accumulation of double-stranded DNA but remain sequence-independent, whereas sequence-specific probes can provide a signal linked more directly to the intended target [16,32]. pH-based reactions require limited buffering capacity because strongly buffered template solutions may suppress the expected color transition, while acidic sample inputs may alter the initial color independently of amplification [30,33].

In a closed-tube format, the indicator is added before incubation and the result is read without reopening the reaction vessel. This approach reduces post-amplification handling and lowers the opportunity for aerosolized amplicons to contaminate subsequent reactions [18,23]. Closed-tube detection does not prevent contamination introduced during blood processing or reagent preparation, and nonspecific indicators cannot distinguish the intended product from primer-derived or non-template amplification [23]. Carry-over control and amplification specificity must therefore be addressed separately. Colorimetric indicators are not interchangeable because their performance depends on both the signal-generating chemistry and the composition of the processed blood input. Representative readout strategies relevant to a closed-tube neonatal-blood workflow are compared in Table 1.

4. Pre-Analytical Challenges and Contamination Control in Neonatal Blood

4.1. Blood Volume, Sampling Probability, and Specimen Timing

The performance of a neonatal blood LAMP assay is determined before amplification begins. Blood volume is the first constraint because neonatal diagnostic testing must be performed from a limited total blood volume [27,37,38]. In a 2024 neonatal intensive care unit (NICU) cohort, the median blood-culture inoculum was 1.0 mL, but 35% of bottles contained less than 0.9 mL [7]. A separate prospective study found that obtaining at least 1 mL was feasible in most neonates, although inadequate volumes were more frequent during late-onset sepsis evaluations in very-low-birth-weight infants [39]. At low microbial densities, detection becomes probabilistic because the tested aliquot may contain no organism even when bacteremia is present [27,40,41]. Experimental work with common neonatal pathogens showed that small inocula were less reliable when concentrations approached or fell below 1 CFU/mL [40]. For molecular testing, the effective input is not the volume collected but the fraction of the original specimen that finally enters the LAMP tube. Every transfer, split, concentration, and elution step should therefore be considered in the analytical sensitivity claim.

Specimen timing and collection source also influence interpretation. Blood should ideally be obtained before antimicrobial administration because culture requires viable organisms, whereas nucleic-acid tests may remain positive after bacterial killing [27,42]. Direct molecular detection can consequently identify microbial DNA when culture is negative, but such discordance does not by itself establish active bloodstream infection [13,27,43]. Collection from an indwelling catheter may increase the recovery of organisms associated with the line while also complicating distinction between catheter colonization, contamination, and systemic infection [44,45,46]. The collection volume, anatomical source, number of attempts, time relative to antibiotics, and interval to processing should be recorded for every clinical sample.

4.2. Pathogen Recovery and Amplification Inhibition

Whole blood is a difficult amplification matrix. Pathogen nucleic acid is present against a large background of human genomic DNA, leukocytes, erythrocytes, plasma proteins, and other substances that can interfere with microbial recovery or downstream enzymatic reactions [27,47,48]. Increasing the input volume may improve the probability of capturing rare organisms, but it also increases host material and inhibitor load [27,48,49,50]. Efficient processing must lyse both Gram-negative and Gram-positive organisms without losing low-copy DNA during purification [51]. This is particularly relevant for organisms with robust cell walls, including staphylococci and streptococci [27]. Recovery from intact organisms in whole blood is therefore dependent on the sample-processing and DNA-extraction method [27,51,52].

Extraction-free processing reduces handling time and consumable use, but it trades purification for dilution or chemical tolerance. A neonatal-blood group B Streptococcus (GBS) LAMP study demonstrated analytical feasibility in spiked specimens, yet spiking does not reproduce the variable cellular composition, antibiotic exposure, pathogen viability, or microbial burden of prospectively collected sepsis samples [19]. Heat treatment or crude lysis must be validated across clinically relevant hematocrits and bacterial concentrations rather than at a single blood dilution. For pH-based colorimetric LAMP, acidic sample components or extraction buffers may shift the starting color, while strongly buffered inputs may suppress the expected endpoint change [30,33]. The template volume must therefore be chosen jointly with the lysis chemistry, reaction buffering, and the selected sample-control architecture.

4.3. Collection Contamination and Amplicon Carry-Over

Contamination can occur during collection, sample preparation, reagent handling, or amplification [53]. Skin commensals are especially problematic in neonates because coagulase-negative staphylococci may represent either true late-onset sepsis or contamination. In a recent neonatal cohort, time to positivity, gestational age, and clinical findings improved discrimination between these possibilities, showing that organism identity alone is insufficient [54]. Umbilical-cord blood can provide a larger sample for early-onset sepsis testing, but its value depends on a rigorously standardized sterile collection procedure [55]. Low-biomass molecular assays also require attention to bacterial DNA present in extraction kits, water, plasticware, or the laboratory environment because background DNA can become visible when the true target concentration is very low [27,56,57].

LAMP creates a large amount of amplicon, making carry-over contamination a persistent risk [23]. Closed-tube color detection reduces the release of products after amplification, but it cannot correct contamination introduced before the cap is closed and does not prevent nonspecific amplification [18,23]. Incorporating deoxyuridine triphosphate (dUTP) into LAMP products and uracil-DNA glycosylase into subsequent reactions can selectively degrade uracil-containing carry-over amplicons before new amplification begins [58,59]. This strategy has reduced false-positive contamination in both bacterial LAMP and colorimetric RT-LAMP formats [59,60]. It does not remove native microbial DNA introduced during specimen preparation, so enzymatic control must complement physical workflow controls.

A neonatal LAMP workflow should separate reagent preparation, specimen processing, and amplification, with unidirectional movement and sealed post-amplification tubes. Each batch should include an extraction blank, no-template control, and target-positive control [61]. A sample-level process control should be introduced before lysis to assess recovery and inhibition [61]. A post-extraction control assesses inhibition only [61]. With sequence-independent color indicators, the control should occupy a separate sealed chamber or use a distinguishable sequence-specific signal, because target and control amplification cannot otherwise be differentiated [62,63]. A target-negative result is valid only when the control succeeds. Control failure should be reported as invalid [62]. Weak or late color changes should be interpreted within the predefined reaction time. Validation reports should state blood volumes, sample source, anticoagulant, storage conditions, extraction recovery, template input, invalid results, and contamination events.

5. Evidence for Neonatal-Sepsis Pathogens

The pathogen content of a neonatal-sepsis assay cannot be defined by a universal list. A recent global meta-analysis found that Gram-negative organisms accounted for 53% of early-onset and 71% of late-onset bacterial sepsis, although dominant species differed markedly by setting [64]. Streptococcus agalactiae was the leading early-onset pathogen overall, whereas Klebsiella species were prominent in both early- and late-onset disease and were especially important in low- and lower-middle-income countries [64]. In a multicenter Southeast Asian dataset, Klebsiella species represented 27.5% of significant isolates, while Candida species accounted for 8.3% of culture-positive episodes [65]. These differences indicate that target selection should follow local neonatal-unit epidemiology rather than the traditional early-onset versus late-onset classification alone.

The most directly relevant evidence concerns Streptococcus agalactiae. An extraction-free cfb-targeted colorimetric LAMP assay demonstrated analytical detection in spiked neonatal blood, supporting matrix feasibility but not clinical diagnostic performance [19]. In a separate neonatal real-time PCR study, detection differed substantially between cerebrospinal fluid and ethylenediaminetetraacetic acid (EDTA) blood, illustrating the importance of specimen-specific clinical validation [66].

Clinical evidence for Escherichia coli has progressed to testing in blood from young infants with suspected sepsis [67]. This study supports the feasibility of rapid LAMP using a low-volume clinical specimen, but neonatal subgroup performance, multicenter reproducibility, and effects on treatment decisions remain uncertain [67]. A pathogen-specific result would still require culture for off-panel organism recovery and phenotypic antimicrobial susceptibility testing [3,13].

Klebsiella pneumoniae LAMP assays have targeted species-identification and hypervirulence markers [68,69], while carbapenemase determinants have been evaluated in separate LAMP reactions [68]. These findings support target selection and modular resistance-marker testing, but they do not establish direct performance in low-volume neonatal blood [68]. Detection of a resistance gene should be interpreted as a genotypic marker rather than a substitute for phenotypic susceptibility testing [68].

Staphylococcal assays demonstrate that species identification and mecA detection can be combined using lateral-flow or compartmentalized microfluidic systems [28,70]. However, neither approach has been validated directly in unprocessed neonatal blood, and only the microfluidic format maintains physical separation during detection [28,70]. For coagulase-negative staphylococci, molecular positivity cannot independently distinguish true bloodstream infection from collection contamination; interpretation still requires collection source, time to positivity, and clinical context [54,71].

Spiked-blood feasibility has been reported for Enterococcus faecalis using LAMP with lateral-flow detection, but direct neonatal-blood validation and a permanently closed colorimetric readout remain absent [72]. Enterococcal targets may be appropriate in locally adapted panels where epidemiology supports their inclusion, but performance should be established in the intended neonatal matrix rather than extrapolated from simulated specimens [64,72].

Fungal LAMP evidence is divided between rapid identification from positive blood-culture bottles and limited direct-specimen evaluation [73,74,75]. Positive-culture assays can shorten species identification after a bottle signals positive, but they do not provide direct-from-blood diagnosis [73]. Direct-specimen evidence remains limited: one study included only two culture-positive whole-blood samples [74], whereas another evaluated spiked healthy-donor blood rather than clinical candidemia specimens [76]. The available evidence therefore differs in both proximity to neonatal clinical use and alignment with a closed-tube colorimetric workflow, as summarized in Table 2.

Across the selected pathogens, the reviewed literature did not identify a prospectively validated, spatially separated panel operating directly from low-volume neonatal blood. Panel composition should remain adaptable to local epidemiology, and a negative panel result must not exclude an off-panel infection [64,65]. Culture should therefore continue for broad organism recovery and phenotypic antimicrobial susceptibility testing [3,13].

6. Clinical Translation and Research Priorities

6.1. Intended Clinical Use

Closed-tube colorimetric LAMP is most appropriately positioned as a rapid adjunct to blood culture during the initial evaluation of suspected neonatal sepsis. Current molecular assays provide faster pathogen detection but show heterogeneous performance and low-certainty evidence, which precludes their use as stand-alone rule-out tests [13]. Culture should proceed in parallel because a target-specific LAMP panel cannot recover off-panel organisms or provide phenotypic antimicrobial susceptibility results [3,13]. The intended clinical contribution is therefore an earlier pathogen-specific result that may support reassessment of empirical therapy while culture remains in progress. A proposed workflow integrating closed-tube colorimetric LAMP with parallel blood culture and clinical interpretation is summarized in Figure 1.

The intended use should be defined before analytical optimization or clinical validation. In hospital settings, the most defensible role is testing symptomatic neonates or infants undergoing a routine sepsis evaluation, rather than universal screening at birth. The World Health Organization (WHO) neonatal-sepsis target product profile and its recent analysis primarily address triage and risk-stratification tests for infants in a diagnostic gray zone [77]. These documents do not directly specify the performance requirements of a pathogen-identification assay, but their emphasis on affordability, accessibility, rapid turnaround, minimal infrastructure, and context-appropriate deployment provides relevant operational guidance [77]. The assay panel and reporting algorithm should consequently be matched to the healthcare level, local pathogen distribution, and the clinical action expected from a positive, negative, or invalid result.

6.2. Diagnostic-Accuracy Validation

The next validation stage should be a prospective multicenter diagnostic-accuracy study enrolling consecutive neonates undergoing routine evaluation for suspected sepsis. LAMP and blood culture specimens should be obtained during the same clinical episode, with collected volume, processed volume, sampling source, and timing relative to antimicrobial administration documented for both methods. Index-test interpretation should be masked to culture and clinical findings, while positivity thresholds, maximum incubation time, and invalid-result criteria should be defined before enrollment. Culture should remain the primary microbiological comparator, but discordant results require cautious interpretation because low inoculated volume and prior antibiotic exposure may reduce culture yield [13,27]. Predefined secondary assessment using clinical findings and independently obtained microbiological evidence may help characterize discordance, but the index-test result should not itself determine the reference classification. Reports should follow the Standards for Reporting Diagnostic Accuracy Studies (STARD) and present participant selection, reference methods, missing specimens, invalid tests, discordant findings, and complete participant flow [78].

6.3. Clinical Utility and Antimicrobial Stewardship

Diagnostic accuracy should be established before clinical utility is tested. A subsequent randomized or pragmatic add-on study should compare standard sepsis management with standard management plus rapid LAMP reporting [79]. Recommended outcomes include time to pathogen-directed therapy, time to antibiotic narrowing or discontinuation, total antibiotic exposure, repeated blood sampling, length of stay, mortality, and cost per clinically useful result [13,80]. Safety outcomes should include missed or delayed treatment of culture-positive and off-panel infections. LAMP-guided decisions should follow a predefined antimicrobial-stewardship algorithm, and empirical treatment should not be withheld when the clinical indication for immediate therapy is strong [81].

6.4. Technical Standardization and Quality Assurance

Technical translation requires a locked assay design, reproducible manufacturing, reagent stability under intended storage conditions, predefined visual or instrument-assisted interpretation, and external quality assessment. Analytical validation should examine strain inclusivity, cross-reactivity, intact-cell detection, clinically relevant organism concentrations, blood-matrix variation, invalid-result frequency, reader agreement, and lot-to-lot performance [15,23]. A compartmentalized panel should report three operational categories: target detected, target not detected with a valid sample control, and invalid because the sample control failed. With any negative panel result, the report should state that an off-panel bloodstream infection is not excluded. The decisive endpoint is therefore not amplification alone, but a reproducible and clinically interpretable result that can be evaluated for its effect on time-sensitive decisions while preserving culture and antimicrobial stewardship.

7. Conclusions

Closed-tube colorimetric LAMP has the potential to provide earlier pathogen-specific information during the evaluation of suspected neonatal sepsis, but its clinical performance will depend on the entire diagnostic pathway rather than amplification speed alone. Low-volume and low-density bacteremia, variable pathogen recovery, blood-derived inhibition, pre-amplification contamination, and nonspecific color changes remain important sources of analytical and interpretive error. The available evidence is derived mainly from isolates, simulated blood matrices, positive blood-culture bottles, and non-neonatal specimens, and does not yet support routine use as a stand-alone diagnostic or rule-out test. Clinical translation will require validated processing of low-volume neonatal blood, compartmentalized pathogen targets, appropriate process and batch controls, predefined result and invalidity criteria, and panel composition adapted to local epidemiology. Blood culture should continue in parallel to detect off-panel organisms and provide phenotypic antimicrobial susceptibility results. The next priority is prospective multicenter diagnostic-accuracy validation in neonates, followed by clinical-utility studies assessing whether rapid reporting can improve antimicrobial decision-making without compromising patient safety.