Click-based determination of accumulation of molecules in Escherichia coli

Domain: neuroscience
Published: 2026-01-24
Editorial outcome: major_revision

Abstract
Gram-negative bacterial pathogens pose a significant challenge in drug development because their outer membranes hinder the permeation of small molecules. The lack of widely adoptable methods for measuring the cytosolic accumulation of compounds in bacterial cells further hinders drug discovery efforts. To address this challenge, we report the development of the Chloroalkane Azide Membrane Permeability (CHAMP) assay, which we designed specifically to assess molecule accumulation in the cytosol of Gram-negative bacteria. The CHAMP analysis utilizes bioorthogonal epitopes anchored within HaloTag-expressing bacteria and measures the cytosolic arrival of azide-bearing test molecules through strain-promoted azide–alkyne cycloaddition. This workflow enables robust and rapid accumulation measurements of thousands of azide-tagged small molecules. Our approach consistently produces comprehensive accumulation profiles that surpass the scale of previous measurements in Escherichia coli (E. coli). We validated the CHAMP assay across various chemical and biological contexts, including hyperporinated cells, membrane-permeabilized cells, and E. coli strains with impaired TolC function, a key component of the efflux pump. The CHAMP platform provides a simple, high-throughput, and accessible method that enables the analysis of over 1000 molecules within hours. This technique addresses a critical gap in antimicrobial research and has the potential to accelerate the development of effective agents against Gram-negative pathogens.

Introduction
The increased prevalence of multidrug-resistant Gram-negative bacterial infections is extremely alarming1. The CDC issued a report in 2019 that categorized bacterial pathogens based on their threat level. The list of pathogens considered as “urgent threats” was primarily populated by antibiotic-resistant Gram-negative bacteria and included E. coli2. Lack of treatment options will usher in the post-antibiotic era, meaning that routine infections can become lethal and standard invasive medical procedures carry a much higher level of risk. Therefore, strategies that aim to address these threats have high significance. We pose, and there is wide agreement in the community3–12, that the lack of robust and widely adoptable tools to measure the accumulation of molecules into bacteria has severely hampered antibiotic drug discovery. The Golden Era of antibiotics leveraged naturally abundant small molecules that were readily identified using traditional methods; since the end of the era, this methodology has proven to be much more difficult to further mine for new antibiotics.

The next phase of antibiotic drug discovery could potentially leverage the wealth of proteomics, genomics, and metabolomics data to design small-molecule agents that are potent and of high specificity. To accomplish this, the field fundamentally needs guiding principles describing the molecular determinants of small molecule permeation into Gram-negative bacterial cells, akin to Lipinski’s rules of 5 (Ro5).

Gram-negative bacterial pathogens pose a significant challenge to antibiotic development due to their distinctive cell wall structure, particularly the outer membrane (OM), which restricts the penetration of small molecules into the periplasmic space (and then further into other subcellular compartments). It is currently understood that small molecules capable of traversing the OM either possess specific physicochemical properties that enable crossing of this barrier by passive diffusion or hijacking porins or other membrane-embedded proteins (Fig. 1a). Typically, minimum inhibitory concentration (MIC) analyses have been used as a proxy for drug accumulation. However, this approach is problematic because MIC reflects antimicrobial activity, which is more indicative of target engagement, rather than directly measuring intracellular concentrations. It is therefore crucial to develop robust and widely adopted live cell assays that can independently measure the accumulation of small molecules across the OM, beyond just relying on MIC values. Developing such methods will provide more accurate insights into the accumulation behavior of potential molecules and ultimately aid in improving antibacterial drug development13.

The field has consistently regarded liquid chromatography-tandem mass spectrometry (LC-MS/MS) as a premier technique for quantifying drug accumulation14–17. Indeed, devoid of the need for a chemical tag, this method presents significant advantages for measuring the uptake of molecules in bacteria. However, mass spectrometry, while advantageous for analyzing whole cell association, has principal limitations as currently reported: (a) restricted throughput capacity and (b) would ambiguously define molecule location unless additional, careful fractionation methods are included to show periplasmic or cytoplasmic accumulation18, and (c) needs compounds to be mass active. This lack of subcellular localization is also a limitation of spectroscopic measurements, such as those using fluorescent tags or inherent fluorescence. Despite these challenges, recent tour de force efforts by the Hergenrother laboratory19,20 have successfully employed LC-MS/MS to develop the eNTRy rules, highlighting the significance of primary amines in the accumulation of small molecules in E. coli. Yet, this level of throughput has been difficult to achieve by others in the field. Consequently, limited throughput in analyzing the cytoplasmic accumulation of molecules remains a significant challenge. Without a clear understanding of the exact parameters that drive the accumulation of molecules into the cytosol of E. coli, efforts to broadly test and prioritize drug leads will be hindered.

In recent years, the Kritzer lab introduced a groundbreaking method for quantifying cytosolic accumulation of molecules in mammalian cells. This method, called the ChloroAlkane Penetration Assay (CAPA), involves applying a chloroalkane-tagged ‘target molecule’ to HaloTag-expressing mammalian cells21,22. The level of cytosolic accumulation is then inferred from the fluorescence signal produced by a chloroalkane-fluorophore ‘chase molecule’. HaloTag, which can be expressed as freely diffusing proteins in the cytoplasm, selectively reacts with chloroalkanes to form a stable covalent bond. Critically, the covalent and subcellular nature of the assay confirms the cytosolic arrival of molecules, rather than whole-cell association23–28. For these reasons, this method is now widely regarded as the gold standard for measuring cytosolic accumulation in mammalian systems. Building on the success of CAPA in mammalian systems, we recently adapted this approach to measure cytosolic accumulation in diderm bacteria, such as E. coli, Mycobacterium smegmatis, and Mycobacterium tuberculosis, in a process we term ‘Bacterial CAPA’ (BaCAPA)29. To address (1) potential interference induced by the 15-atom chloroalkane tag in bacterial accumulation studies (Figs. 1b) and (2) limitations in the availability of chloroalkane-tagged test molecules, we developed an enhanced method in E. coli that substitutes the chloroalkane with a shorter 3-atom azide tag. The choice of the azide tag was based on its minimal size, relatively low impact on the physicochemical properties of the parent compound, and stability in biological systems30,31. Moreover, there are well-established protocols for late-stage azide installation into complex molecules32,33 and in synthetic peptides, as well as metabolic incorporation within biomacromolecules, such as RNA, oligosaccharides, and proteins34–37.

Our team recently introduced the Peptidoglycan Accessibility Click-Mediated Assessment (PAC-MAN) assay in mycobacteria38. In PAC-MAN, a strained alkyne, dibenzocyclooctyne (DBCO), is metabolically incorporated39–41 into the peptidoglycan layer of diderm bacteria to monitor the passage of molecules through the OM. In this work, we adapted our previous workflow by treating HaloTag-expressing E. coli cells with a chloroalkane-DBCO reagent that anchors a strained alkyne to cytosolic HaloTag. Azide-tagged test compounds that reach the bacterial cytosol are then covalently imprinted to HaloTag via strain-promoted azide-alkyne cycloaddition (SPAAC)42 reactions (Fig. 1c). This step is followed by treatment with an azide-tagged fluorophore, after which cellular fluorescence is measured using a flow cytometer or a plate reader in 96- or 384-well plates. Fluorescence levels are inversely proportional to the extent of SPAAC reactions between DBCO and the test azides in the cytosol, providing a straightforward method to determine apparent cytoplasmic arrival (Fig. 1d). The irreversible covalent reaction between chloroalkane-DBCO and HaloTag anchors DBCO in the cytosol and offers a method for determining subcellular localization. This adaptation, termed the ‘CHloroalkane Azide Membrane Penetration’ (CHAMP) assay, focuses on measuring cytosolic accumulation rather than whole-cell association. It operates at a throughput level that, to our knowledge, has not been previously achieved on such a scale. We demonstrate the capability to make more than 5000 measurements of approximately 1500 unique molecules in various biological contexts.

Results
We first optimized the assay parameters to maximize the dynamic range of the fluorescence signals and adapted the assay for compatibility with high-throughput screening platforms. A small panel of azide-tagged molecules was used to benchmark the testing phase, followed by testing a series of antibiotics with varying levels of biological activity against E. coli. Next, this approach was validated by evaluating the impact of OM permeabilization in E. coli on the accumulation of azide-modified drugs and analyzing the reactivity of these azides with DBCO-modified beads. Furthermore, the assay was adapted for high-throughput screening platforms, with the datasets coupled with structural modifications that were previously proposed to impact accumulation in E. coli. We analyzed a highly diverse set of azide-tagged molecules, including a 404-member and a 1152-member library, to evaluate their cytosolic uptake in E. coli under four unique biological contexts: wildtype, tolC-null E. coli (the major efflux pump gene), porin overexpression (highly permeable/hyperporinated), and OM permeabilization induced by antibiotic treatment. These thousands of unique measurements, all acquired in under a month, were analyzed to deliver a first-in-class profiling platform in E. coli.

Optimization of HaloTag Expression in E. coli
Optimizing key conditions in the CHAMP assay is essential before using it as a method for assessing molecule accumulation in bacterial cells. These conditions include HaloTag expression, anchoring of DBCO-chloroalkane onto HaloTag, choice of fluorophore, and conditions for measuring the accumulation of the test molecules. First, E. coli cells were transformed with a plasmid encoding HaloTag (see SI for details). The expression of functional HaloTag was confirmed by treating cells with a chloroalkane-linked rhodamine 110 (R110cl) and subsequently conducting flow cytometry analysis (Fig. 2a). Given the nature of the dye, it was expected to have a high level of cellular accumulation. The observed cellular fluorescence intensity correlated well with the concentrations of IsoPropyl β-D-1-ThioGalactopyranoside (IPTG) used during the protein expression phase (Fig. 2b). Of significance, the cellular fluorescence intensities initiated at a low value in the absence of IPTG, suggesting that the expression of HaloTag was responsible for capturing the chloroalkane-linked fluorophore in the cytosolic space of E. coli. The signal intensities exhibited exponential growth with increasing IPTG concentration before reaching a saturation plateau.

Additionally, with the goal of optimizing the concentration of the anchoring chloroalkane ligands in the CHAMP assay, cells expressing high levels of HaloTag were subjected to varying concentrations of R110cl. Cellular fluorescence intensities were low in the absence of R110cl, while they experienced significant enhancement at 0.3 μM of R110cl, followed by a dose-dependent increase that plateaued at approximately 3 μM (Fig. 2c). The localization of the fluorescence signals within the cells was additionally assessed using confocal imaging (Figs. 2d and S1). To confirm that the accumulation measurements in this study reflect cytoplasmic localization of molecules, the cytoplasmic compartment of E. coli cells was delineated by labeling the cell wall with D-Lys(FITC)-OH, which labels the peptidoglycan layer43. Consequently, the cytosolic probe for confocal imaging was changed to coumarin chloroalkane (Comcl), which fluoresces in a distinct spectral window, thereby enabling clear spatial separation of cytosolic and periplasmic signals. Moreover, fluorescence imaging of an SDS-PAGE gel containing cellular components from induced cells revealed a fluorescent band at approximately 30 kDa, consistent with the expected molecular weight of HaloTag (Fig. 2e). Together, these results confirm the capability of HaloTag to selectively anchor molecules tagged with a chloroalkane linker in E. coli.

DBCO-chloroalkane anchoring within HaloTag
To mitigate the potential impact of a longer chloroalkane linker, we employed an azide tag that could then undergo an in cyto SPAAC reaction30,31 with DBCO. To accomplish this, we envisioned that we could anchor a DBCO epitope within HaloTag by pre-treating cells with DBCOcl (Fig. 2f). HaloTag-bound DBCO can then be leveraged to assess the accumulation of azide-bearing target molecules in the cytosol of E. coli. Several experiments were conducted to refine the procedure. First, E. coli cells expressing HaloTag were incubated with DBCOcl to install the DBCO reporter within the cytoplasm. Empirical assessments were conducted to determine the optimal concentration for DBCOcl anchoring, employing an approach reminiscent of BaCAPA experiments. Cells were incubated with varying concentrations of DBCOcl followed by the treatment with R110cl (Fig. S2). Our findings indicated that the maximum levels of DBCO installation were achieved at approximately 50 μM of DBCOcl, exhibiting an EC50 value of ~18.5 μM.

Subsequently, we conducted a baseline CHAMP experiment by treating E. coli cells with the azide-tagged rhodamine 110 (R110az) to reveal the anchored DBCO handles within individual cells. A titration of R110az revealed a concentration-dependent increase in fluorescence intensities, establishing the optimal concentration of R110az for subsequent experiments (Fig. 2g). Satisfyingly, a large dynamic range in fluorescence of up to 30-fold was observed in cells treated with both DBCOcl and R110az compared to controls. In the absence of DBCOcl treatment, cellular fluorescence levels were similar to those of untreated cells, indicating that R110az has minimal retention in bacterial cells without a covalent SPAAC reaction. Cellular fluorescence levels were also near background levels in non-induced cells that were also treated with DBCOcl and R110az, suggesting negligible retention of DBCOcl or cytosolic DBCO modified with R110az (Fig. S3).

Confocal microscopy confirmed that the fluorescence signal was distributed throughout the cytoplasm (Fig. 2h and S4) and fluorescent gel imaging also showed a band matching in the expected molecular weight range of HaloTag (Fig. 2i). These results are consistent with a site-selective anchoring of the DBCO and subsequent SPAAC with an azido-fluorophore at the site of HaloTag. When HaloTag was expressed in a smooth E. coli strain (ATCC 25922 with intact lipopolysaccharides on its outer surface), there was a similar dynamic range (Fig. S5). These data highlight the potential ability to translate CHAMP to a system with more complex surface composition and suggest that CHAMP can be applied more broadly across most types of bacteria that can be genetically manipulated. In the case of smooth E. coli, it is important to consider how the system can work in this context because surface biomacromolecules (including O-antigen) can alter accumulation profiles due to the potential impact of O-antigens in steric shielding of porins44,45. To verify the viability of the E. coli cells after inducing HaloTag expression and subjecting cells to DBCOcl treatment, we conducted a SYTOX Green analysis46. In live prokaryotic cells, SYTOX Green is excluded from cells with intact permeability barriers47. Our results revealed that the expression of HaloTag and incubation of these cells with DBCOcl and its occupancy by HaloTag in E. coli did not compromise the integrity of the cytoplasmic membrane (Figure S6). This finding emphasizes the suitability of utilizing CHAMP for investigating molecular permeation to the cytosol without causing disruption to the cell’s permeability barriers.

We next sought to assess the compatibility of CHAMP with other azide-bearing fluorophores. Specifically, two more azide-bearing fluorophores, fluorescein (Flaz) and coumarin (Comaz), were tested. Incubation of cells after DBCOcl treatment with either fluorophore resulted in a significant increase in the cellular fluorescence (Fig. 2j). Cells incubated with Flaz exhibited a considerably lower fluorescence intensity compared to R110az-treated cells. Nevertheless, those results confirm the versatility of the assay for using various azide-bearing fluorophores. Recognizing that the assay is effectively completed at the end of the incubation with the test molecule, we proposed that higher signal-to-noise ratios could be achieved by removing the permeation barrier to the azido-fluorophores. Notably, formaldehyde fixation has been previously shown to outperform alcohol-based fixation in terms of increasing permeability in E. coli48. Our results indicated that fixation performed prior to the incubation with fluorophores led to an overall increase in fluorescence levels for all three tested molecules (Fig. S7). This observation suggests that the dyes themselves could face significant permeation challenges imposed by the intact envelope of E. coli. Among the three dyes, the most favorable signal-to-noise ratio in fixed cells was achieved with Flaz treatment, resulting in a substantial increase in cellular fluorescence with minimum background staining.

Influence of OM permeabilization on accumulation of Cy3az and Cy5az
We then applied CHAMP to analyze the impact of OM permeabilizers. Of significance, the permeation of small molecules in Gram-negative bacteria is purported to be primarily impeded by the OM49–52. This formidable barrier has been a focal point of previous investigations aimed to identify compounds capable of disrupting the OM integrity. These types of molecules can serve as antibiotic adjuvants to enhance the effectiveness of antibiotics or circumvent resistance mechanisms through co-treatment. A prominent example in this category is polymyxin B nonapeptide (PMBN)53, a nonapeptide fragment derived from polymyxin B (Fig. 2k). PMBN is of particular interest due to its ability to permeabilize the OM at low concentrations. In our hands, PMBN displayed no significant impact on the colony-forming unit (CFU) count when administered at concentrations of 5 μM and 10 μM, consistent with previous studies demonstrating its low toxicity at these concentrations (Fig. S8)54.

To analyze the impact of OM permeabilization by PMBN on the accumulation of molecules, we selected the sulfonated cyanine series as model compounds. These dyes contain an azide group, specifically Cy5az and Cy3az, whose permeabilities are limited by their sulfonate groups (Fig. 2l, m). We hypothesized that OM permeabilization by PMBN would enhance the accumulation of the dyes, thereby increasing cellular fluorescence. Briefly, live E. coli cells were either pre-exposed to low concentrations of PMBN or co-incubated with PMBN and dye to assess the accumulation of Cy5az and Cy3az. Our data revealed a striking 19-fold increase in cellular fluorescence when PMBN was pre-incubated with the cells, and an even more substantial 34-fold increase when the cells were co-incubated with PMBN and Cy5az (Fig. 2l). A full range of permeabilization by PMBN was observed by 2.5 μM with an EC50 of 0.8 μM (Fig. S9). This pattern was mirrored with the analogous but smaller fluorophore, Cy3az, signifying that OM permeabilization assisted the accumulation of slightly less hydrophobic dyes as well (Fig. 2m). Given the substantial dynamic range that was observed, we propose that this assay could also be adapted for detection using a microplate reader for high-throughput screening. Indeed, our results showed that the dynamic range was mostly retained in this format (Fig. S10), which should provide a more direct path to screening campaigns with diverse molecular libraries for OM permeabilizers.

Accumulation of test small molecules and antibiotics
We set out to establish the feasibility of CHAMP in monitoring the accumulation of azide-tagged molecules into E. coli cells. For this initial demonstration, E. coli cells with DBCOcl anchored on expressed HaloTag were titrated with a test azide-bearing molecule (Fig. 3a), followed by treatment with R110az, and cellular fluorescence levels were measured via flow cytometry. A concentration-dependent reduction in cellular fluorescence was observed, revealing an EC50 of ~14 μM. The fluorescence intensity reached a maximum value at approximately 50 μM of the test molecule. These findings confirm the capability of the optimized CHAMP assay for investigating the accumulation of molecules in E. coli. With these results in hand, we shifted our focus to testing a series of azide-tagged small molecules featuring diverse structural motifs, anticipating that these variations would influence their accumulation. In the first subset, 1–3, we tested the potential impact of amidation of carboxylic acids (Fig. 3b). Indeed, we observed that carboxamide, 1p, had a higher level of apparent accumulation relative to its carboxylic acid counterpart 1n (Fig. 3c). Appreciating the potential for an internal hydrogen bond due to the amino group in the β-position of C-terminal 2,4-diaminobutyric acid and the potentially confounding behavior of 1p/1n, we extended the amino group further from the β-carbon in 2n/2p. The apparent accumulation level of 2p was significantly more pronounced, which may reflect the greater contribution of the positively charged lysine side chain, as it cannot readily form an internal hydrogen bond with the C-terminal group. While compounds 1n/1p/2n/2p are theoretically expected to be primarily positively charged or net neutral under physiological pH, we sought to test if the pattern would hold true for a similar compound lacking a basic amino group. Our results showed that carboxamide in 3p still led to higher levels of cytosolic accumulation in E. coli relative to the net negative 3n. This small panel of compounds demonstrates how specific structural variations can be readily tested for their potential ability to drive accumulation in E. coli, which could be crucial in designing prodrugs that mask carboxylic acids to enhance their permeability in similar pathogenic bacteria.

The Hergenrother group has used an established method in the field, mass spectrometry, to empirically demonstrate that primary amines can serve as privileged functional groups to improve accumulation in E. coli as part of the eNTRy rules17,19,20. To benchmark E. coli CHAMP with these structural parameters, we made three additional compound panels, 4–6 (Fig. 3d). Series 4 has a similar general scaffold to our established molecules (3) that exhibited high levels of accumulation, while the molecules (4a-4e) feature five different configurations of the side chains ranging from an unfunctionalized terminal hydrocarbon to amino groups that vary in their degrees of methylation. Within this series, it was observed that the substitution of a methyl group in 4a for a primary amino group in 4b resulted in a large increase in apparent accumulation (Fig. 3e). The alkylation of the amino group progressively reduced accumulation levels in 4b-4e. These results show marked agreement with the eNTRy rules. In fact, the trimethyl 4e had a similar accumulation profile as the compound 4a, which does not have an amino group. To test the generalizability, a subsequent series, 5, was synthesized that retained many of the same features to test the generalizability. Interestingly, the hydrocarbon-terminated 5a displayed higher apparent accumulation than any other analog with an amino group, regardless of methylation states.

Next, we pivoted to benchmark the assay using an alternative strategy. Instead of using a pulse-chase modality to measure apparent accumulation, the test compounds in 6 all have a small fluorophore incorporated as part of the molecule (Fig. 3f). The goal was to determine whether the pulse-chase method produces results consistent with those obtained from a single-step direct fluorescence analysis. Similar to the 4 and 5 series, the methylation of the amino groups in series 6 led to lower accumulation levels (Fig. 3g). The absence of DBCOcl that installed the landmark and IPTG led to near background levels of fluorescence, which suggests that the cellular fluorescence observed in series 6 is due to engagement with the landmark and reported on cytosolic arrival (Fig. S11). Within the same series, we reasoned that we could evaluate the CHAMP-based levels of accumulation using a pulse-chase workflow. Consistently, we observed that the pattern of accumulation across 6a-6e was nearly identical for a direct measurement and as a CHAMP-based analysis with a non-overlapping fluorophore (Fig. S12). Together, these findings align well with the eNTRy rules and underscore the advantages of having a robust and straightforward method for measuring cytosolic accumulation. Such a method can significantly enhance the throughput of identifying structural determinants of accumulation.

We posited that CHAMP would serve as a versatile platform for unraveling the molecular determinants of accumulation related to elements found within the OM, which can include both the membrane bilayer and the proteins embedded within this barrier. To achieve this, we sought to compare the apparent accumulation of azide-tagged antibiotics (Fig. 4a). Our first goal was to evaluate the accumulation of azide-tagged antibiotics across biological contexts related to efflux pumps (Fig. 4b)55. Efflux pumps can greatly reduce the concentrations of antibiotics by excreting them to the extracellular space. Of the 36 known or putative efflux pumps in E. coli56, AcrAB-TolC is the best characterized efflux pump and has been described as the most prevalent in decreasing susceptibility to antibiotics57,58. Genetic deletion of tolC is sufficient to disrupt the tripartite formation of the AcrAB-TolC complex59,60. To test CHAMP in this context, we adopted an antibiotic that has been previously described as a canonical substrate of AcrAB-TolC. Novobiocin belongs to the class of aminocoumarin antibiotics, and it inhibits bacterial DNA gyrase61. Given its cytosolic target, novobiocin needs to permeate past the OM in order to reach the target gyrase. Deletion and mutations to tolC can dramatically alter the MIC of novobiocin, which has been ascribed to its substrate recognition by this powerful efflux pump62.

An azide-modified novobiocin derivative, novAZ1, that minimally altered the chemical configuration of the original antibiotic (Fig. 4a). In WT E. coli, CHAMP analysis revealed that novAZ1 had a low level of apparent accumulation relative to tolC-null E. coli (Fig. 4c), which is an indication that TolC could reduce the effective concentration of novAZ1 in the cytosol of WT E. coli. These results are consistent with novAZ1 being a substrate of TolC. A parallel strategy was also employed to test the viability of CHAMP in measuring substrate recognition in E. coli. It is well established that the function of TolC can be disrupted with small molecule inhibitors such as phenylalanine-arginine b-naphthylamide (PAβN)63. To this end, a CHAMP analysis was performed for novAZ1 with and without the co-incubation of PAβN. Our results showed that the use of a small molecule chemical inhibitor of TolC led to an elevated level of apparent accumulation of novAZ1. Collectively, these experiments validate novAZ1 as a substrate of AcrAB-TolC in the context of the CHAMP assay. In the future, we will explore how CHAMP can be paired with high-throughput screens to discover next-generation TolC inhibitors (or those of other efflux pumps).

With these results in hand, we set out to widen our library of azide-tagged antibiotics and measure their apparent accumulation in E. coli in four different contexts that operate at the OM. The antibiotics covered a wider range of mechanisms of action and cellular targets. The azide groups were added via late-stage modifications, and the chemistry used varied depending on the functional groups available to perform edit to install the azide epitope. In all, 11 azide-tagged antibiotics were included in this subpanel, representing one of the most comprehensive analyses reported to date. Analysis of the accumulation profiles revealed that CHAMP was readily able to discern the relative cytosolic accumulation levels of a wide range of molecules. Of note, there was no detectable accumulation of vancomycin azide (vanAZ1) within the cytosolic space of E. coli, consistent with expectations for this organism. Vancomycin exerts its activity by binding to the D-Ala-D-Ala terminus of the lipid II peptidoglycan precursor, which is presented on the outer leaflet of the inner membrane64. However, E. coli, and, more broadly, Gram-negative bacteria are intrinsically resistant to vancomycin due to the impermeability of their outer membrane, and only becomes susceptible when this barrier is disrupted65. Nonetheless, the CHAMP assay, as employed in this study, provides information that excludes cytoplasmic accumulation of vancomycin but remains agnostic to its potential presence in the periplasm. Several antibiotics act within the periplasmic space of E. coli and other Gram-negative bacteria, including β-lactam antibiotics, which target periplasmic penicillin-binding proteins (PBPs) involved in peptidoglycan synthesis, and glycopeptide antibiotics such as vancomycin, which inhibit cell wall cross-linking66,67. The current inability of the CHAMP platform to discriminate periplasmic localization represents a limitation of the assay in its present form. Accordingly, vancomycin serves as an effective negative control in our CHAMP assay for assessing the intracellular accumulation of other antibiotics. Other molecules that were smaller in size (e.g., trimethoprim, triAZ1, and sulfonamides, sulAZ1) showed effective accumulation into the cytosol of E. coli. Among molecules with high accumulation levels were oxazolidinones. A similar profile was also observed for smooth E. coli that has its full set of O-antigen (Fig. 4d), which indicates that the O-antigen chains do not appear to significantly hinder the accumulation levels of these specific molecules.

Next, accumulation analysis via CHAMP was performed across the entire series of antibiotics using either the TolC-null E. coli strain or the small molecule AcrAB-TolC inhibitor. Interestingly, a number of antibiotic derivatives, including puromycin, showed differential accumulation in tolC-deleted strains relative to the parental WT strain (Fig. 4e). For some of these types of structures, the differential accumulation in tolC-null strains had yet to be analyzed. Additionally, accumulation profiles for the antibiotic derivatives were also tested with WT cells treated with the small molecule inhibitor, PAβN. As expected, the profile of accumulation across the series of antibiotics was not identical (Fig. 4f). This difference can be attributed to many factors, including the disruption to the OM in the absence of TolC and/or the polypharmacology of most small molecules, including PAβN. Testing a broader range of antibiotics could start to reveal how cells respond to genetic or small-molecule stressors in the context of drug accumulation in E. coli. These results demonstrate the potential of utilizing CHAMP to confirm inhibitors of the TolC efflux pump and, most importantly, offer a mean to readily compare the apparent accumulation of antibiotic derivatives in strains with genetic alterations that can potentially modulate the accumulation of drugs.

Another aspect of the OM that CHAMP could investigate is how modifying its overall integrity affects the accumulation of modified antibiotics. This includes OM-embedded channels that facilitate passage across the OM and the chemical permeabilization of this critical bilayer. We projected that for molecules whose accumulation was impeded by the OM, there would be an increase in apparent accumulation after the passage was improved. As described before, PMBN was chosen as a chemical permeabilizer of the OM. Critically, a high number of antibiotics had a shifted apparent accumulation in E. coli cells upon the co-treatment with PMBN (Fig. 4g). These results clearly demonstrate that the treatment with OM permeabilizers can alter the level of otherwise less permeable molecules into E. coli. Alternatively, we tested a genetic strategy to increase the accumulation levels in E. coli in a strain with overexpression of the porin FhuA68. Our CHAMP results using the antibiotics panel clearly show that, for some antibiotics, there is a shift in their accumulation profile upon hyperporination. Interestingly, the two methods of improving passage across the OM were distinct and driven by their modes of action. Whereas PMBN may have a broader impact, and, theoretically, also impact the cytoplasmic membrane integrity69, the substrate scope of FhuA is likely to be much narrower because it is a porin. In this work, an open and non-selective variant of FhuA that does not discriminate based on hydrophilicity, designated “pore” was used; the pore is localized to the outer membrane. The substrate size range for this strain is not clear, but E. coli K12 BW25113 expressing this variant of FhuA was shown to be sensitive to vancomycin with a 16-fold reduction in MIC relative to the wild type strain70. Nonetheless, these analyses provide an extensive description of the changing landscape of accumulation in four biological contexts that can impact passage and retention within bacteria. To the best of our knowledge, this is one of the most comprehensive analyses of a series of antibiotic-derivatives within biological contexts that could impact accumulation.

High-throughput library analyses in E. coli
Our investigation shifted to evaluating larger sets of molecules in E. coli. To this end, we assembled a library of 404 commercially available azide-bearing compounds (see SI). This collection encompassed a broad spectrum of characteristics, including hydrophobicity, rigidity, hydrogen bond donors/acceptors, molecular weight, charge, and polarity. Recognizing the prevalence of azide groups in various steric and electronic configurations, we pondered whether these chemical attributes could influence their reactivity with DBCO. To gauge the reaction progress of DBCO-azide within the stipulated assay timeframe, we empirically assessed the reaction completion employing DBCO-modified beads (Fig. S13). A workflow akin to live cell analysis was executed, albeit devoid of the permeability barriers inherent to the cellular outer and inner membranes. Interestingly, it was discerned that not all azides exhibited satisfactory reactivity within the designated timespan of the assay, signifying that azide reactivity necessitates careful consideration (Fig. S14). It is noteworthy that previous investigations have also documented variations in reactivity profiles between azides and strained alkynes, including DBCO71. Aromatic azides have been found to have lower reactivity in SPAAC reactions, with nearly a sevenfold reduced reactivity of phenyl azide in comparison with aliphatic azides, even where the aliphatic azide conformation is in the form of azidotoluene. As for alkyl azides, the following trend in reactivity has been observed: primary > secondary > tertiary. This statement is supported by prior studies72. It was quickly recognized that variations in the intrinsic reactivities of azides would constrain the analysis to only those azides exhibiting comparable reactivity, thereby narrowing the chemical diversity and reducing the overall screening breadth achievable with the CHAMP platform.

In light of the challenges posed by the variable reactivities of azides from a large library that are outsourced, we embarked on an alternative approach. To mitigate the influence of azide reactivity, we employed a differential scan, examining the same set of molecules in two closely related biological contexts that exhibit phenotypic divergence. This allowed us to isolate the change in apparent accumulation and attribute it solely to the underlying phenotypic differences. We posited that this method would be apt for unraveling the molecular determinants imposed by the OM of E. coli. To illustrate this, our first test case was the analysis of the accumulation of a 404-molecule library in WT E. coli with and without the treatment with PMBN (Fig. 5a). As before, the apparent accumulation of small molecules, for which the OM served as the primary barrier to entry into the cytosol, is expected to change upon co-incubation with PMBN, as this barrier will be disrupted. Conversely, molecules with high propensity for accumulation in intact WT E. coli cells would be expected to have minimal differences in their apparent accumulation with PMBN treatment. From our analysis, there was a consistent shift in apparent accumulation across the majority of molecules tested upon the co-incubation with PMBN (Fig. 5b). These results demonstrate the significant barrier that the OM poses to cytosolic arrival and highlight the feasibility of conducting large-scale analyses across various biological contexts. CHAMP can effectively inform us about the differences in the behavior of molecules within these contexts. Further, the difference in the accumulation profile was analyzed across a range of physicochemical parameters, including hydrogen bonding numbers, rotatable bonds, molecular weight, topological polar surface area, ClogP, and logS (Fig. 5c). A larger increase in apparent accumulation following PMBN treatment should theoretically indicate that a specific parameter is implicated in reducing accumulation to the cytosol. These results highlight how high-volume screening can rapidly reveal patterns of molecular features that influence cytosolic accumulation. In this demonstration, more than 800 individual measurements were made (>400 in wild-type and >400 in PMBN-treated cells), making it the largest reported screen in E. coli to date.

As a complement to using a site-selective structural modification series (such as the aminomethylation), the late-stage tagging of antibiotics, and the 404 commercial library of azides, we next set out to evaluate the accumulation profile of a larger non-commercial library of small molecules. Each molecule in this library is azide-tagged, making it compatible with CHAMP31. These molecules are diverse: aromatic and aliphatic, covering a wide range of chemical space and spanning a range of molecular weights. Our objective was to thoroughly examine the feasibility of a large-scale scan that has not been reported to this point. To further demonstrate the scalability of CHAMP, we screened a library comprising 1152 azide-bearing compounds that was recently described by the Dong and Sharpless groups in their strategy to install azide tags onto molecules using a diazotizing reagent32. A benchmarking analysis was performed, spanning a range of cellular fluorescence levels: high, low, and intermediate, using a 96-well plate format. Our experiments affirmed the remarkable reproducibility of CHAMP, underscored by a Z’ score of 0.954 (Fig. S15).

The entire 1152-member library was subjected to bead analysis under the same conditions as the cellular assay. Our results showed that there was a range of reactivities that were observed within the timescale of this assay, owing to the differences in the environments of the azide groups in a large library (Fig. S16). To start, WT E. coli cells were subjected to E. coli CHAMP analysis, and the results were compared with the reactivity on the beads (Fig. 6a). In this analysis, there is a marked shift in the plot toward the upper left quadrant of the plot. This shift is highly indicative that their engagement with the DBCO landmark is greatly improved upon the removal of the physical barrier (e.g., OM and cytosolic membrane). Subsequently, we applied CHAMP to the complete array of the large library of molecules in four biological contexts: WT E. coli, WT E. coli pre-incubated with PMBN, hyperporinated E. coli, and hyperporinated ΔtolC E. coli strains. Our objective was to empirically demonstrate that CHAMP has a throughput capability that is unlike any technology to date, and, when paired with a large library, it provides a workflow to rapidly analyze cytosolic accumulation in E. coli. As with the smaller set of molecules, there was a general shift in apparent accumulation in WT E. coli upon the pre-treatment with PMBN (Fig. 6b). These results demonstrate that chemical disruption to the membrane barriers can broadly improve the cytosolic residency of molecules across a diverse range of physicochemical properties. Hyperporinated E. coli could provide a passage point to molecules, as in the case of many antibiotics. In comparison to WT E. coli, there was a general increase in apparent accumulation of molecules in the hyperporinated E. coli strain (Fig. 6c). Interestingly, there was general agreement on the two modalities of reduced barriers (Fig. 6d), yet not all compounds behaved the same way. This deviation could be, in part, due to the nature of substrate recognition by porins. Alternatively, the potential for the cytosolic membrane disruption by PMBN could reveal a slightly different set of molecules that arrive in the cytosol. With the goal of leveraging CHAMP to more broadly assess the substrate recognition by TolC, we compared the library response in hyperporinated E. coli to hyperporinated ΔtolC E. coli (Fig. 6e). This was an attempt to reveal compounds that are TolC substrates but have no access to the sites of recognition due to a lack of accumulation. In this comparison, there was surprisingly general agreement in the behavior of the molecule in the presence and absence of TolC, which could indicate that the reduced barrier in hyperporinated cells could overwhelm the efflux capacity of TolC.

Using the comparison between WT and ΔtolC (Fig. 6f), we sought to broadly explore how the structure of the molecule could impact TolC recognition. We examined the correlation between compound efflux and physicochemical properties (Fig. S17). A few properties are shown to correlate with high statistical significance, including hydrogen bond acceptor, conformational flexibility (number of rotatable bonds), amide bond, polar surface area, polarizability, saturated carbon atoms, larger molecular size, and less globularity, which correlate with a higher chance of chemicals being recognized by TolC. There was a large range of chemical structures whose accumulation profiles were found to be especially altered by the deletion of tolC (Fig. S18). As part of the effort to understand how much reactivity weighs in on the analysis, we performed correlation analysis with different reactivity cutoff values to filter out compounds with low reactivity. Surprisingly, the reactivity cutoff from 0 to 0.4 does not have a significant influence on correlation values. We theorize that there may be two principal factors at play. First, by performing the differential scan of wild-type and tolC deletion mutant, the contribution from reactivity is minimized. Secondly, while the reactivity may still amplify the difference between the wild-type and the ΔtolC mutant, the poorer reactivity means a smaller difference in the differential scan. For this reason, such data points make smaller contributions to the final correlation valued during the correlation calculation.

Next, we set out to identify motifs that correlate with compound efflux, with the understanding that this would help extract sub-structure feature that make a compound more likely to be recognized by TolC. We initially employed the Bemis-Murcko scaffold method to identify specific scaffolds whose accumulation profiles were particularly sensitive to the presence of TolC. The Bemis-Murcko scaffold is a method that treats rings and the connection between the rings (linkers) as the scaffold. The remaining fragments of the molecule are then considered as side chains. This library of 1152 compounds contains 158 unique scaffolds in total, with some of them much more enriched, e.g., the top 2 scaffolds have more than 10 compounds in each scaffold (Fig. 6g). We selected the first 73 scaffolds having more than 1 member in each scaffold and analyzed the average accumulation shift upon tolC deletion. We used a cutoff of 0.05 and depicted scaffolds where the average accumulation shift is more than 0.05. Such scaffolds include piperidine, pyrrolidine, piperazine, thiophene, naphthalene, azetidine, etc. Piperazine-based compounds have been shown to be potential modulators of efflux pump AcrAB-TolC73,74. Thiophene antibacterials are known to be substrates of AcrAB-TolC in E. coli75. Aryl piperidines have been screened as potentiators of novobiocin activity in E. coli cells by high-throughput assays76.

Though Murcko scaffolding represents the skeleton of a molecule, it is possible that side chains may also play an important role in substrate recognition by the TolC-related efflux pumps that are not captured in the scaffold-based analysis. Therefore, we also performed a chemical clustering analysis of the library. Chemical clustering is an approach that finds similarities among the molecules and groups them by similarity. This is also a common approach77 in drug discovery campaigns to prioritize compounds, select representative molecules, and derive smaller datasets. In our case, we grouped the molecules and investigated whether specific chemical clusters have a larger accumulation shift upon tolC deletion. We distributed the library into 24 clusters through hierarchical clustering. We then performed hierarchical clustering. The accumulation shift profile for molecules in each cluster showed two clusters (6 and 19) that have significantly higher apparent accumulation in the absence of TolC (Fig. 6h). Cluster 6 contains many compounds with the carbamate and ester functional group, while cluster 19 contains doxorubicin and a similar derivative. We further developed a hypothesis that the carbamate motif may contribute to compound efflux, though it is not clear if the carbamate or the ester group alone is enough for that. To validate this hypothesis, we calculated accumulation shift upon tolC deletion for combinations of these functional groups (Fig. 6i). Only carbamate-based compounds were more populated within this dataset. Together, these results highlight the types of analyses that can be conducted and the queries that can be parameterized using a large volume of accumulation profiles in pathology-relevant biological contexts.

Methods
Biological methods
Plasmid construction for HaloTag expression in E. coli. For expression of the HaloTag protein in WT E. coli, (smooth, ATCC 25922) and in different variants of E. coli K12 (BW25113), HaloTag was cloned onto an expression vector pQE-30 (Genscript) and codon-optimized for expression in E. coli; the molecular cloning and DNA synthesis were performed by Genscript (Table 1).

HaloTag protein amino acid sequence. MASAEIGTGFPFDPHYVEVLGERMHYVDVGPRDGTPVLFLHGNPTSSYVWRNIIPHVAPTHRCIAPDLIGMGKSDKPDLGYFFDDHVRFMDAFIEALGLEEVVLVIHDWGSALGFHWAKRNPERVKGIAFMEFIRPIPTWDEWPEFARETFQAFRTTDVGRKLIIDQNVFIEGTLPMGVVRPLTEVEMDHYREPFLNPVDREPLWRFPNELPIAGEPANIVALVEEYMDWLHQSPVPKLLFWGTPGVLIPPAEAARLAKSLPNCKAVDIGPGLNLLQEDNPDLIGSEIARWLSTLEISGGGGGSGGGGSGGGGSGS

HaloTag protein expression and bacterial cell culture. For expression of the HaloTag protein in WT E. coli, (smooth, ATCC 25922) and in different variants of E. coli K12 (BW25113), the expression plasmid Halo_pQE-30 was transformed into Escherichia coli and grown on an LB/agar plate with ampicillin (100 μg/mL) at 37 °C overnight. Colonies were picked and grown first in 5 mL LB broth with ampicillin at 37 °C overnight. A flask containing 35 mL of LB media was then inoculated with the overnight culture at a ratio of 1:100 in the presence of 100 μg/mL ampicillin and cells were grown at 37 °C for 2 h, or until the optical density at 600 nm reached 0.2 when cultures were induced with 1 mM Isopropyl β-d-1-thiogalactopyranoside (IPTG) at 37 °C for 2 h in a shaker incubator. Cells were then pelleted at 3005 xg for 3 min. The pellet was washed three times with 1X PBS pH 7.4, following which cells were resuspended in 35 mL of 1X PBS for the assay. The expression and identity of the protein were confirmed by SDS-PAGE. Uncropped and unprocessed scans of the SDS-PAGE gel can be found in the Source Data master EXCEL file.

One-step assay with rhodamine chloroalkane (R110cl). 3 mL of HaloTag-expressing cell culture was transferred to 15 mL conical tubes and pelleted by centrifuging. The cells were washed 3X with 3 mL of 1x PBS and finally re-suspended in 3 mL of 1x PBS. The cells were pipetted into wells in a 96-well plate in triplicate, pelleted down by centrifuging for 3 min at 2700 xg, and resuspended using 5 μM R110cl. The cells were incubated for 30 min at 37 °C, centrifuged for 3 min, washed 1X with PBS, and fixed with 4% formaldehyde in PBS for 30 min statically at room temperature. The cells were subjected to analysis by flow cytometry on the Attune NxT Acoustic Focusing Cytometer (Invitrogen) by exciting using the blue laser with emission at 525 nm for acquisition.

CHAMP assay development
Three-step assay: - DBCOcl incubation, azido-tagged small molecule (pulse), and Flaz dye (chase). 35 mL of HaloTag expressing fresh cell culture was washed with PBS and transferred to 50 mL conical tubes and pelleted by centrifuging. The cells were re-suspended in 35 mL of a 50 μM solution of DBCOcl in PBS and transferred to an Erlenmeyer flask for incubation at 37 °C for 30 min in a shaker incubator. The DBCOcl imprinted cells were centrifuged to pellet at 3000 xg using an HERAEUS Multicentrifuge X1 centrifuge (Thermo fisher Scientific), washed 3X with PBS, and resuspended in the same volume of PBS as the original cell suspension. 90 μL of this cell culture was added to each well of a pre-prepared 96-well plate containing 10 μL of the azido-tagged test compound from each 500 μM working solution in PBS. The plate was then covered with plastic film and incubated for 1 h at 37 °C in a shaker incubator. Each azido-tagged test molecule was replicated in three wells. Control experiment wells were either on the same plate or a separate plate in the same shaker incubator for 1 h at 37 °C, and each well contained 10 μL of PBS with 90 μL of DBCO-labeled cell culture (both IPTG-induced and non-induced cells), in triplicate. In the end, the plate was then spun at 2700 xg for 3 min using a plate holder centrifuge to pellet the cells, the supernatant was discarded, and the cells were washed 1X with PBS. Next, cells in both the experimental wells and the control wells were fixed with 4% formaldehyde in PBS for 30 min statically and incubated without shaking at room temperature on the benchtop. Following formaldehyde fixation, the cells were centrifuged and washed 1X with PBS. In the last chase step, 100 μL of 50 μM Flaz was added to the washed cells in each well, resuspended by pipetting up and down, and incubated at 37 °C for 60 min in a shaker incubator. Finally, the cells in the plate were pelleted by centrifuging in a plate centrifuge, washed 1X with PBS, resuspended in 200 μL of PBS, and subjected to analysis by flow cytometry.

Two-step assay without fixation prior to chase step: - DBCOcl incubation, azido-tagged dye. DBCO-labeled cells were prepared as described earlier and resuspended in the same volume of PBS as the original cell suspension. 90 μL of the cell culture was added into each well containing 10 μL of the azido-tagged dye at 500 μM in 96-well plates, covered with plastic film, and incubated for 1 h at 37 °C in a shaker incubator. Each azido-tagged molecule or dye was replicated in three wells. The plate was then spun at 2700 xg for 3 min using a plate holder centrifuge, the supernatant was discarded, and the cells were washed 1X with PBS. Next, cells were fixed with 4% formaldehyde in PBS for 30 min statically at room temperature on the benchtop. Following formaldehyde fixation, the cells were washed 1X with PBS and resuspended in 200 μL of PBS for analysis by flow cytometry.

Two-step assay with fixation prior to chase step: - DBCOcl incubation, formaldehyde fixation, and azido-tagged dye. DBCO-labeled cells prepared as described earlier were washed 3X with PBS and resuspended in 4% formaldehyde in PBS and incubated statically at room temperature on the benchtop for 30 min. Following formaldehyde fixation, the cells were washed 1X with PBS and resuspended in the same volume of PBS as was in the original cell suspension. For the azido-tagged dyes screen, 90 μL of the cell culture was added into wells containing 10 μL of the azido-tagged dye at 500 μM in 96-well plates, covered with plastic film, and incubated for 1 h at 37 °C in a shaker incubator. Each azido-tagged dye test was replicated in three wells. The plate was then spun at 2700 xg for 3 min, the supernatant was discarded, and the cells were resuspended in 200 μL of PBS for analysis by flow cytometry.

Assay with PMBN: - DBCOcl incubation, azido-tagged dye co-incubation with PMBN
DBCO-labeled cells were prepared as described earlier and washed 3X with PBS and resuspended in PBS containing 5 μM PMBN. For the azido-tagged small molecule (three-step) or azido-tagged dyes screen, 90 μL of the cell culture was added into each well containing 10 μL of the azido-tagged small molecule/dye at 500 μM in 96-well plates, covered with plastic film, and incubated for 1 h at 37 °C in a shaker incubator. Each azido-tagged molecule or dye was replicated in three wells. The plate was then spun at 2700 xg for 3 min using a plate holder centrifuge, the supernatant was discarded, and the cells were washed 1X with PBS for fixing, as described earlier, before analysis by flow cytometry (two-step), or for the chase step with an azido-tagged dye (three-step), as described earlier.

1152 azido-tagged test molecules library screen
The 1152-molecule azido-tagged library in the form of 96-well mother plates (total 12 plates) was obtained from the Dong Lab at the Institute of Translational Medicine, Zhangjiang Institute for Advanced Study (Shanghai Jiao Tong University) and stored at -20 oC. Each well contained 50 μL of individual test molecules at 5 mM in DMSO. Prior to the assay, the original 96-well plates (denoted as the “mother plates”) were allowed to warm up to room temperature and spun down for 1 min at 2700 xg, to allow the stock solution to settle at the bottom of each well, using a plate holder centrifuge. 10 μL of the DMSO stock was drawn from each well and dispensed to a new set of plates (denoted as the “daughter plates”). Then 90 μL PBS was added to the daughter plates to make a 100 μL solution of each test molecule at 500 μM working concentration. 10 μL of each test molecule from the daughter plates was transferred to a new 96-well plate (the “working plates”).

DBCO-labeled cells were prepared as described earlier and poured into a reservoir. 90 μL of DBCO-labeled cell suspension was drawn from the reservoir and dispensed to the working plates prepared above using a 96-multichannel pipette dispenser (Sequence®). The plates were covered with plastic films and incubated for 1 h at 37 °C in a shaker incubator. The plates were then spun at 2700 xg for 3 min using a plate holder centrifuge. The supernatant was discarded, and the cells were washed 1X with PBS. Next, cells were fixed with 4% formaldehyde in PBS for 30 min statically at room temperature on the benchtop. Following formaldehyde fixation, the cells were washed 1X with PBS. For the final chase step, 100 μL of 50 μM Flaz was added to each well, resuspended, and incubated at 37 °C for 1 h in a shaker incubator. Lastly, the cells were pelleted by centrifuging the 96-well plate in a plate holder centrifuge, washed 1X with PBS, resuspended in 200 μL of PBS, and analyzed by flow cytometry.

Cell preparation for confocal microscopy
DBCO-labeled cells prepared as described earlier were washed 3X with PBS and resuspended in the same volume of PBS as was drawn from the original cell suspension. For the azido-tagged small molecules screen, 90 μL of the cell culture was added into wells containing 10 μL of the azido-tagged test compound at 500 μM in 96-well plates, covered with parafilm, and incubated for 1 h at 37 °C in a shaker incubator. Each azido-tagged dye test was replicated in three wells. Control experiment wells were either on the same plate or a separate plate in the same incubator for 1 h at 37 °C, and each well contained 10 μL of PBS with 90 μL of DBCO-labeled cell culture (both IPTG-induced and non-induced cells), in triplicate. The plate was then spun at 2700 xg for 3 min in a plate holder centrifuge, the supernatant was discarded, and the cells were washed 1X with 1X PBS.

Next, cells in both the experimental wells and the control wells were fixed with 4% formaldehyde in 1X PBS for 30 min statically incubated at room temperature on the benchtop. Following formaldehyde fixation, the cells were washed 1X with PBS, and for the chase step, 100 μL of 50 μM Flaz was added into each well, resuspended by pipetting up and down, and incubated at 37 °C for 60 min in a shaker incubator. Lastly, the cells were pelleted by centrifuging the 96-well plate in a plate centrifuge, washed 1X with PBS, and resuspended in 200 μL of PBS. For confocal microscopy, the fixed cells were treated with 5 μg/mL of TAMRA-tagged wheat germ agglutinin (Vector Laboratories, RL-1022) for 30 min at 4 °C. Glass microscope slides were spotted with a 1% agar pad, and 5 μL of the sample was deposited onto the agar. Samples were covered with a glass coverslip and imaged using a Zeiss 880/990 multiphoton Airyscan microscopy system (40× oil-immersion lens) equipped with 488 nm and 550 nm lasers. Images were obtained and analyzed via Zeiss Zen software. We acknowledge the Keck Center for cellular imaging and the usage of the Zeiss 880/980 multiphoton Airyscan microscopy system (PI-AP: NIH-OD025156).

Confocal fluorescence images of E. coli cells expressing HaloTag, labeled with D-Lys(FITC), and treated with coumarin chloroalkane to produce a blue cytosolic signal. E. coli cells harboring the plasmid pQE-30_HaloTag (Amp) were cultured overnight in LB medium. The overnight culture was diluted 1:100 into fresh LB medium supplemented with 500 µM D-Lys (FITC) and incubated at 37 °C with shaking until the optical density at 600 nm (OD₆₀₀) reached approximately 1.2 to allow the incorporation of D-Lys(FITC) in the cell wall. Protein expression was induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and cells were incubated for 1 h to allow expression of the HaloTag protein. Uninduced cultures were maintained as negative controls. Next, cells were washed three times with phosphate-buffered saline (PBS) and incubated with 50 µM coumarin chloroalkane (Comcl) for 30 min at 37 °C with shaking. Cells were subsequently washed three times with PBS, fixed with 4% (w/v) formaldehyde for 30 min, washed again, and mounted on 1% (w/v) agarose pads for imaging by confocal microscopy. Confocal fluorescence imaging was performed using a Leica STELLARIS 8 Super-Resolution Microscope equipped with a 100× oil-immersion objective. Image acquisition and analysis were carried out using ImageJ software. Excitation was performed using 405 nm (Coumarin) and 488 nm (FITC) laser lines. We acknowledge the Keck Center for cellular imaging (PI-AP: NIH-OD025156).

Confocal fluorescence images of E. coli cells expressing HaloTag, labeled with D-Lys(FITC), and treated with DBCOcl, which was subsequently reacted with Comaz to generate a blue cytosolic signal. E. coli cells harboring the plasmid pQE-30_HaloTag (Amp) were cultured overnight in LB medium. The overnight culture was diluted 1:100 into fresh LB medium supplemented with 500 µM D-Lys (FITC) and incubated at 37 °C with shaking until the optical density at 600 nm (OD₆₀₀) reached approximately 1.2 to allow the incorporation of D-Lys(FITC) in the cell wall. Protein expression was induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and cells were incubated for 1 h to allow expression of the HaloTag protein. Uninduced cultures were maintained as negative controls. Next, cells were washed three times with phosphate-buffered saline (PBS) and incubated with 50 µM DBCOcl for 30 min at 37 °C with shaking. Cells were subsequently washed three times with PBS, fixed with 4% (w/v) formaldehyde for 30 min, washed again, and incubated with 50 µM Comaz for 1 h at 37 °C with shaking. Finally, cells were washed three times and mounted on 1% (w/v) agarose pads for imaging by confocal microscopy. Confocal fluorescence imaging was performed using a Leica STELLARIS 8 Super-Resolution Microscope equipped with a 100× oil-immersion objective. Image acquisition and analysis were carried out using ImageJ software. Excitation was performed using 405 nm (Coumarin) and 488 nm (FITC) laser lines. We acknowledge the Keck Center for cellular imaging (PI-AP: NIH-OD025156).

Novobiocin azide (novAZ1) accumulation assay in WT E. coli BW25113 and ΔtolC E. coli BW25113 with or without efflux inhibitor PAβN. DBCO-labeled cells for the accumulation assay were prepared as described earlier. For each strain of E. coli, cells were divided into two equal groups, with one set containing 25 μL of 200 μM novAZ1 and 25 μL of PBS as a control, and the other set containing 25 μL of 200 μM novAZ1 and 25 μL of 64 μg/mL PAβN. Next, 50 μL of DBCO-labeled WT E. coli cell suspension or ΔtolC E. coli suspension was added to the wells in volumes needed to bring the final concentration of novAZ1 and PAβN to 50 μM and 16 μg/mL, respectively. These cells were incubated for 1 h, then pelleted by centrifugation, and the supernatant was removed. Cells were washed 3X in PBS and resuspended for the chase step with R110az as described earlier. Finally, the cells were centrifuged, washed 1X with PBS, fixed with 4% formaldehyde statically at room temperature, and subjected to flow cytometry analysis. A similar method was followed for all azido-tagged antibiotics used in this study.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "Summary of what the authors did:\nIn this manuscript entitled “Click-Based Determination of Accumulation of Molecules in Escherichia coli” the authors\ndeveloped the Chloroalkane Azide Membrane Penetration (CHAMP) assay that quantifies the accumulation of molecules in\nthe cytoplasm of Escherichia coli. Previous studies have estimated accumulation of a molecule in the bacteria through the\nminimal inhibitory concentration (MIC). However, the MIC reflects the molecule’s ability to inhibit a molecular target, as\nopposed to its accumulation in the cytoplasm.\nCHAMP is a combination of the Peptidoglycan Accessibility Click-Mediated AssessmeNt (PAC-MAN) and the Chloroalkane\nPenetration Assay (CAPA). Bacteria expressing HaloTag are treated with a chloroalkane-tagged strained alkyne\n(dibenzocyclooctyne; DBCO), anchoring DBCO onto HaloTag. The molecule of interest, modified with a small azide handle,\nis incubated with the cells and if it reaches the cytoplasm, it will undergo strain-promoted azide-alkyne cycloaddition\n(SPAAC) with the DBCO-modified HaloTag. Subsequently, bacteria are chased with an azide-tagged fluorescent molecule\nto capture unoccupied HaloTag-DBCO molecules. Similar to CAPA the fluorescent reading is inversely proportional to the\npenetration of the molecule of interest.\nThe authors present careful optimization of CHAMP in E. coli. They show that this assay can be applied to individual\nmolecules with small alterations to their structures to create a structure-accumulation relationship (SAR). Next, they apply\nCHAMP to natural product antibiotics (such as puromycin and vancomycin) and large libraries of molecules (consisting of\n404 and 1152 molecules), highlighting the high-throughput nature of CHAMP.\nThrough CHAMP the authors showed the ability to enhance the accumulation of certain molecules by increasing\npermeability of the outer membrane and inhibiting the activity of the efflux pump TolC. Careful structure-accumulation\nanalysis highlighted particular structural features that enhanced accumulation and export by TolC. Taken together, CHAMP\ncould be important in driving future antimicrobial development studies by rapidly “weeding out” poor candidates that are\nunable to accumulate in the bacterial cytoplasm.\nGeneral comments\nCHAMP is a powerful assay for measuring the accumulation of small molecules in bacteria that has the potential to become\na “guiding force” in future antimicrobial design efforts. Importantly the power of the assay stems from being high-throughput,\nallowing for a large number of reads to be made quickly.\nThe structure and use of language in the manuscript has led to a clear story with minimal errors. A very strong feature of this\nmanuscript is the extensive meta-analysis of their data, linking the structural features of their molecular library to the efflux\nexporter TolC and the accumulation of the molecules of interest in the cytoplasm. Moreover, the data presented here is a\nstrong indicator of the potential of antimicrobial cocktails, coupling TolC inhibitors with antibiotics that otherwise do not\naccumulate as well in E. coli.\nIt is important to note, however, that CHAMP is a “spin-off” of CAPA applying many of its important features (such as\nHaloTag, proximity ligation and chasing with a fluorophore) to a bacterial system, as opposed to a mammalian one. Also,\nCHAMP has few differences compared to their recent Bacterial CAPA (BaCAPA) methodology, essentially switching the\nchemical handle on the molecule of interest and adding chloroalkane-DBCO on HaloTag in E. coli. The authors have done\na good job acknowledging this.\nIn light of this being a thorough and well-written manuscript, only minor additional experiments and corrections have been\nprovided below. If they are addressed in a satisfactory manner, I believe this manuscript would make a great addition to the\nimpactful papers published in Nature Communications.\nPoint-by-point comments:\nIntroduction (page 4) – The authors should make a stronger case about how there is a big problem with antimicrobial\ndevelopment at the beginning of the introduction and not wait until the discussion to bring this up.\nFigure 1 - Very well made. Easy to understand the premise of this assay.\nFigure 2d and (page 9) – For clarity, the authors should show an image of the cells without treatment with R110cl and\nwithout IPTG treatment to establish the background. This is important as the gels in 2e and 2i suggest leaky expression.\nFigure 2e – Authors should appreciate and acknowledge that there is some “leaky” expression of HaloTag (in the absence\nof IPTG) as shown in lane 3 of the fluorescent gel in figure 2e. It would be more thorough to have an additional lane in that\ngel where they transform the cells with the same vector lacking the HaloTag gene (that should show no expression) and\ncarry out R110cl treatment to show that there is no “off-target effect”.\nFigure 2h – Similar to 2d, to account for leaky expression it would be useful to show an image of the cells that have not\ntreated with DBCOcl, R110cl or IPTG.\nFigure 2j – In the left plot, there is also some indication of leaky expression of HaloTag.\nThis is also reflected in Figure S8b and Figure S9.\nFigure 3a and page 14 – On page 14 the authors state that for the test molecule, there is “an EC50 of approximately 3 µM”.\nLooking at figure 3a, it looks like the EC50 should be higher, closer to 10 or 11 µM. Should be double checked.\nFigure 3a – Since in CHAMP the signal is inversely proportional to the accumulation, it important to mention, in the text\nand/or figure legend, how the depicted plot was derived.\nFigure 5 – For the last panel, HBA (hydrogen bond acceptor) and HBD (hydrogen bond acceptor) should be defined in the\nlegend for easy comprehension.\nFigure 6 – Excellent analysis.\nFigure 6i – In the legend of 6i it says “Permeability shift for compounds with or without (left) carbamate group, (middle) ester\ngroup, (right) carbamate”. There seems to be a mismatch between what is shown on the figure and what is described in the\nlegend. The left panel in the figure is labelled as “amide” not carbamate.\nDiscussion, page 32 – “…are porin imbedded within the OM” should be “…is porins imbedded within the OM”.\nDefine replicates: In figures where experiments were carried out in replicates (such as figures 2, 3, 4 and many\nsupplementary figures), it is important to state if the replicates were technical or biological in the legend of the corresponding\nfigure.", "reviewer_2": "This manuscript uses HaloTag as a route to anchor a cyclooctyne inside E. coli. Then entry of azide-linked small molecules\ncan be quantified by their competition for reaction of cyclooctyne with a membrane-permeable azide-fluorophore. Entry is\nquantified for a huge library of different azides, along with analysis in E. coli bearing different alterations to its outer\nmembrane.\nI found this study innovative and the experiments thoroughly take forward the concept.\nI thought that the data were strong and showed careful statistical analysis. There is one issue to address there: the\nmicroscopy in Fig.2 is not informative without showing alongside equivalent negative control images.\nI think it is an excellent example of the use of click chemistry in cells to address a problem of medical relevance. I think it will\nbe of high interest to a wide range of scientists.\nThe authors do a good job in considering the possible effect of the differential reactivity of azides with DBCO. The authors\nshould comment in the Discussion on other possibilities for read-outs from the assay that do not precisely relate to the entry\nof the small molecule-azide of interest:\nPerhaps-\n1.the small molecule azide gets modified by the cell to some other compound\n-some molecule with different entry/exit properties\n-the azide could be reduced, so blocking reaction with DBCO\n2.the small molecule azide alters thiol levels in the cell which changes the non-specific reactivity of the cyclooctyne\n3.rapid lytic activity of the small molecule-azide\n4.Engagement of the small molecule-azide with macromolecular targets hindering DBCO reaction\n5.Potential impact of azide-bearing impurities in each small molecule-azide stock which may have higher permeability.\nThis is not a criticism of the approach but every assay has the potential for hits with alternative explanations, so it is good to\ncontemplate such possibilities early on and consider how to account for them.\nMinor points:\nEven with a magnifying glass it is hard to see what is happening to the HaloTag in cells in Fig. 1d. It would be nice to edit\nthe figure so this part is easier for readers to pick up.\nAlso, it is not ideal that the fluorescence seems to be accumulating in Fig. 1D on the outside of the cell, which could lead to\nconfusion.\nFig. 2l,m It would help readers to draw Cy5az and Cy3az in their most likely charge state\nFig. 5c Give units for surface area\nFig. 6g: Murcko and not Murko\nFig. 6i legend has carbamate twice\n“Interestingly, it was discerned that not all azides exhibited satisfactory reactivity within the\ndesignated timespan of the assay, signifying that azide reactivity necessitates careful\nconsideration (Fig. S12). It is noteworthy that previous investigations have also\ndocumented variations in reactivity profiles between azides and strained alkynes,\nincluding DBCO.55”\nNice to have one or two sentences to summarize which kinds of azides react badly.", "reviewer_3": "Ongwae et al present a new method to study molecule uptake into E. coli. The paper is an impressive and extensive piece of\nwork that addresses an important scientific problem (how to quantify molecule uptake) and that is of relevance to a critical\nsocietal problem (antibiotic resistance). In principle, I feel positively about this manuscript as I think the authors did a\nthorough job in developing and testing their assay and I believe that it will be of good use to the scientific community.\nHowever, I have noticed a number of inaccuracies and mistakes, as well as some open questions in the paper and I would\nlike to see these corrected/addressed before endorsing publication.\n1. The authors several times embellish the limitations of alternative methodologies and in some cases the statements are\nplainly wrong. The authors' work should stand for itself without putting other approaches down and wrong facts should\ncertainly not be used as arguments for the superiority of their method. Some examples:\nPage 4/5: Subcellular localization can be assessed with MS, if proper cell fractionation protocols with sufficient washing\nsteps are followed. The same applies to fluorescent tags and inherent fluorescence, if used spectroscopically. Even more,\nthe latter can be used in microscopy approaches, which, in most cases, easily allows assessment of subcellular localization.\nPage 9: There simply is no challenge to distinguish membrane from cytosol in bacteria. The difference between cell surface\nassociation, membrane localization, and cytosolic localization are glaringly obvious, even with simple wide-field\nmicroscopes. Even the outer and inner membrane can be easily distinguished from each other when following appropriate\nprotocols. The authors even show this themselves with abundantly clear cytosolic fluorescence of their probe.\nPlease check through the manuscript and remove faulty claims.\n2. The authors describe the SYTOX green assay wrong (page 11). Firstly, SYTOX dyes are perfectly cell wall-permeable.\nThe cell wall is the peptidoglycan layer, not the whole cell envelope structure built up of inner membrane, peptidoglycan cell\nwall, and outer membrane. In order to penetrate, SYTOX dyes need both the outer and inner membrane compromised. Thus,\nthey indicate disruption of membrane (not cell wall) integrity. Further, since the inner membrane needs large pores to allow\nthe passage of the dye, which will kill the cells, SYTOX dyes indicate dead cells. So what the authors have proven here is\nsimply that Halo tag expression does not simultaneously create large holes in the inner and outer membrane of E. coli.\nPlease rephrase this part to correct this.\nA second issue with this assay is that it simply does not support the authors' conclusions. Because SYTOX dyes need both\nthe inner and outer membrane compromised, a negative result cannot exclude that either one of them may be damaged. If\nthe authors truly want to make the claim that the cell envelope as a whole is not damaged, they must assess inner and outer\nmembrane integrity separately. This could, for example, be achieved using NPN for the outer and DiSC(3)5 for the inner\nmembrane.\n3. The authors misrepresent/misinterpret vancomycin. On page 18, they say that no intracellular accumulation of vancomycin\nwas observed, which would be consistent with the lack of biological activity against E. coli. Vancomycin binds to lipid II,\nprecisely to the D-Ala-D-Ala motif of the pentapeptide chain. This is a cell surface structure that is presented on the outer\nleaflet of the inner membrane. Naturally, vancomycin would never be found intracellularly, not even in sensitive bacteria.\nHowever, vancomycin is perfectly able to bind to lipid II and inhibit E. coli, if the outer membrane is compromised. The\nauthors even show this themselves (page 19). The authors further imply (by lumping vancomycin in with puromycin and\nothers) that what they observe is uptake into the intracellular space. As outlined above, that is not the case with vancomycin,\nwhich binds to the inner membrane.\nPlease correct these errors and also generally pay more attention to distinguishing cell and cell envelope structures and\ndefining uptake appropriately (as in uptake into the periplasm or cytoplasm) as this is of crucial relevance to the content of\nthe paper.\n4. In connection with that, the authors (wrongly) remark that microscopy would pose difficulties in distinguishing the\nmembrane from the cytosol, but their method clearly cannot achieve this as outlined above. I would be really curious to see\nmicroscopy assays here. For example, vancomycin and puromycin uptake in WT compared to delta tolC would be a good\nsetup to show (i) whether their method os also be well-suitable for microscopy (which would be really exciting in my opinion)\nand (ii) whether it actually can distinguish subcellular localization.\n5. Since the Halo tag is expressed in the cytosol, I am wondering how it could detect vancomycin uptake (which sits in the\nmembrane as outlined above). I can imagine four scenarios: (i) vancomycin flips over the membrane to the inner leaflet and\nis then able to interact, (ii) the Halo tag is somehow also expressed in the periplasm where it would encounter vancomycin,\n(iii) the azide label alters vancomycin properties enabling cytosolic translocation, or (iv) the experimental conditions do\ncompromise the inner membrane so vancomycin and/or Halo tag can cross this barrier. Each one of these would have rather\ncrucial implications for the usability of the method and I would strongly suggest to address this.\n6. Somewhat related to the same question, I would be really curious if periplasmic localization would be possible to see.\nWould the system work with periplasmic expression?\n7. The authors make some mistakes describing PMBN. Firstly, they keep talking about sublethal concentrations (page\n12/13), which implies that PMBN is lethal. However, it has no activity against E. coli (under standard laboratory conditions)\nthat would show in standardized MIC assays (MICs usually above 512 µg/mL). Secondly, they state that it could interfere\nwith the inner membrane (page 20). However, that is not the case (see for example PMID 36165741). The reference they cite\nfor this does not state this either. Please correct this.\n8. Also regarding this, on page 20 where the authors argue with the possible inner membrane effects of PMBN, it would\nmake much more sense to argue that FhuA has a lower substrate range because it is a porin, which is naturally limited in the\nsize and charge of molecules that it lets pass. PMBN induces large-scale destruction of the OM, letting much larger\nmolecules through including massive molecules like nisin.\n9. On page 28 the authors argue with using MIC changes as proxy for uptake. While the argument that MICs depend on\ntarget interaction is of course true, it is also weak in the context of the paper as the authors argue that structural changes to\ncompounds will affect target interaction. However, that is also the case for the azide linkers used here. The argument could\nfurther be broken down to testing the activity of compounds with and without PMBN or in WT and a permeable mutant,\nwhich is the same setup the authors are using here and hence not really that different as a concept.\nSimilarly, on page 30/31 the authors argue for the usability of their method in screening for potentiators that disrupt outer\nmembrane integrity. While I do not doubt that this could an application, the argument is also weakened by the observation\nthat their assay is itself impacted by the integrity of the outer membrane, making such measurements hard to calibrate.\n10. Related to the last point, I would wish that the authors would more honestly and critically reflect on the limitations of their\nmethod and in an open and honest way compare the pros and cons between their method and existing ones. The\nembellishment of the limitations of competing techniques and the absence of honest reflections on the limitations of theirs is\na red thread in the paper that I cannot condone.", "reviewer_4": "Antibiotics must overcome the cell wall and accumulate at the target site. In particular for Gram-negatives this is a real\nchallenge with no simple technique to quantify the uptake. As the authors point out correctly there is a true need for simple\nmethods allowing to quantify accumulation. Mass spectrometry has seen a huge gain in sensitivity allowing larger\nthroughputs. Here the authors suggest a novel technique to quantify the uptake of a particular class of antibiotics. This\nmethod has the potential for high throughput. The work is carefully done with a broad range of proofs and tests. I recommend\nstrongly publication.\nBelow are a few comments:\n- The manuscript is difficult to read for outsiders. First, I suggest a more basic scheme about the function bringing the subject\nto a broader audience.\n- What is the sensitivity, how many bacteria are needed? Could this extend to single cells?\n- I congratulate the authors for the amount of work and test. This is a very good basis for other works. However, I read the ms\nseveral times to catch the bigger picture. Maybe restructuring might render the ms easier to read? (I understand that its not\nthe primary goal to make it easy).\n- Please clarify the limits of the approach."}, "author_response": "Comment1:\nIntroduction (page 4) – The authors should make a stronger case about how there is a\nbig problem with antimicrobial development at the beginning of the introduction and not\nwait until the discussion to bring this up.\nResponse: Thank you so much and sorry for this confusion; the description of the\nproblem of antimicrobial development has been moved back to the introduction – page\n4.\nComment 2:\nFigure 1 - Very well made. Easy to understand the premise of this essay.\nResponse: Thank you very much for the kind feedback. We're glad to hear that the\ndesign and clarity of Fig.1 effectively conveyed the assay's premise.\nComment 3:\nFigure 2d and (page 9) – For clarity, the authors should show an image of the cells\nwithout treatment with R110cl and without IPTG treatment to establish the background.\nThis is important as the gels in 2e and 2i suggest leaky expression.\nResponse: We thank the reviewer for this advice. Fig. 2d and Fig. S1 have been updated\nto include labeling of the cell wall using D-Lys(FITC)-OH (green), thereby delineating the\ncell envelope (Angew Chem Int Ed Engl. 2012 Dec 7;51(50):12519-23. doi:\n10.1002/anie.201206749). Consequently, the cytosolic label in this image was switched\nto coumarin chloroalkane (Comcl), which fluoresces in a distinct spectral window,\nenabling clear differentiation of the cytosol from the periplasmic space. Additionally, E.\ncoli cells not induced with IPTG and treated with Comcl have been included as a negative\ncontrol to establish the baseline fluorescence background.\nFig. S1. |\nFig. 2d and Fig. S1. | Confocal fluorescence images of E. coli cells expressing\nHaloTag, labeled with D-Lys(FITC) and treated with coumarin chloroalkane to\nproduce a blue cytosolic signal. E. coli cells harboring the plasmid pQE-30_HaloTag\n(Amp) were cultured overnight in LB medium. The overnight culture was diluted 1:100\ninto fresh LB medium supplemented with 500 µM D-Lys(FITC)-OH and incubated at 37\n°C with shaking until the optical density at 600 nm (OD ) reached approximately 1.2 to\nallow the incorporation of D-Lys(FITC)-OH in the cell wall. Protein expression was\n₆₀₀\ninduced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and cells were\n2\nincubated for 1h to allow expression of the HaloTag protein. Uninduced cultures were\nmaintained as negative controls. Next, cells were washed three times with phosphate-\nbuffered saline (PBS) and incubated with 50 µM of coumarin chloroalkane (Comcl shown\non panel a) for 30 min at 37 °C with shaking. Cells were subsequently washed three times\nwith PBS, fixed with 4% (w/v) formaldehyde for 30 min, washed again and mounted on\n1% (w/v) agarose pads for imaging by confocal microscopy. Confocal fluorescence\nimaging was performed using a Leica STELLARIS 8 Super-Resolution Microscope\nequipped with a 100× oil-immersion objective. Image acquisition and analysis were\ncarried out using ImageJ software. Excitation was performed using 405nm (Coumarin)\nand 488 nm (FITC) laser lines; panel b shows the images in different channels. Scale bar:\n2 µm.\nComment 4:\nFigure 2e – Authors should appreciate and acknowledge that there is some “leaky”\nexpression of HaloTag (in the absence of IPTG) as shown in lane 3 of the fluorescent\ngel in figure 2e. It would be more thorough to have an additional lane in that gel where\nthey transform the cells with the same vector lacking the HaloTag gene (that should\nshow no expression) and carry out R110cl treatment to show that there is no “off-target\neffect”.\nResponse: We thank the reviewer for this advice. Fig. 2e has been revised to include\nadditional lanes representing cells harboring the empty vector, as well as cells cultured in\nthe absence or presence of IPTG and treated with or without R110cl. These additions\nprovide a more comprehensive comparison of HaloTag expression and labeling under the\nvarious experimental conditions.\nlabeling conditions. Coomassie-stained and fluorescence-imaged gels showing HaloTag\nprotein expression and labeling efficiency in E. coli cells carrying the HaloTag-expressing\nplasmid. Cells were analyzed with or without IPTG induction and in the presence or\nabsence of R110cl treatment to evaluate protein expression and labeling performance.\nComment 5:\nFigure 2h – Similar to 2d, to account for leaky expression it would be useful to show an\nimage of the cells that have not treated with DBCOcl, R110cl or IPTG.\nResponse: We thank the reviewer for this helpful suggestion. Fig. 2h and Fig. S4 have\nbeen updated to include E. coli cells that were not induced with IPTG but treated with\ncoumarin azide to establish the baseline fluorescence background. The use of coumarin\nazide was necessary because D-Lys(FITC)-OH (green) was employed to label the cell\nwall, thereby delineating the cell envelope (Angew Chem Int Ed Engl. 2012 Dec\n7;51(50):12519-23. doi: 10.1002/anie.201206749). Consequently, a cytosolic probe\nemitting in a distinct spectral window was required to clearly distinguish the cytosol from\nthe periplasmic space.\nFig. S4. |\nwith D-Lys(FITC) and treated with DBCOCl, which subsequently reacted with\nComaz to generate a blue cytosolic signal. E. coli cells harboring the plasmid pQE-\n30_HaloTag (Amp) were cultured overnight in LB medium. The overnight culture was\ndiluted 1:100 into fresh LB medium supplemented with 500 µM D-Lys (FITC)-OH and\nincubated at 37 °C with shaking until the optical density at 600 nm (OD ) reached\napproximately 1.2 to allow the incorporation of D-Lys (FITC)-OH in the cell wall. Protein\n₆₀₀\nexpression was induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and\ncells were incubated for 1h to allow expression of the HaloTag protein. Uninduced cultures\nwere maintained as negative controls. Next, cells were washed three times with\nphosphate-buffered saline (PBS) and incubated with 50 µM DBCOcl for 30 min at 37 °C\nwith shaking. Cells were subsequently washed three times with PBS, fixed with 4% (w/v)\nformaldehyde for 30 min, washed again, and incubated with 50 µM Comaz (on panel a)\nfor 1h at 37 °C with shaking. Finally, cells were washed three times and mounted on 1%\n(w/v) agarose pads for imaging by confocal microscopy Confocal fluorescence imaging\nwas performed using a Leica STELLARIS 8 Super-Resolution Microscope equipped with\na 100× oil-immersion objective. Image acquisition and analysis were carried out using\nImageJ software. Excitation was performed using 405nm (Coumarin) and 488 nm (FITC)\nlaser lines; panel b shows the images in different channels. Scale bar: 2 µm.\nComment 6:\nFigure 2j – In the left plot, there is also some indication of leaky expression of HaloTag.\nThis is also reflected in Figure S8b and Figure S9.\nResponse: We thank the reviewer for taking note of this. We observed a modest degree\nof leaky expression of the HaloTag protein from this plasmid, which contributes to the\nbaseline fluorescence detected in our measurements. Notably, the absolute magnitude of\nthis background signal is dye dependent. As shown in Fig. 2j (left panel), the dye Flaz\nexhibits relatively high background fluorescence and non-specific binding compared with\nother dyes. Although Flaz displays limited accumulation within E. coli cells, it is likely\nassociated with cell surface components, resulting in measurable background even after\n6\nstandard 3× washing. Consequently, when fluorescence from uninduced (–IPTG) cells is\nnormalized to that of induced (+IPTG) cells, the low intracellular accumulation of Flaz\nyields a reduced signal-to-noise ratio, which can exaggerate the apparent effect of leaky\nexpression. In contrast, as shown in the right panel, the dye Comaz exhibits low\nbackground and accumulates more efficiently in E. coli, resulting in a higher signal-to-\nnoise ratio and minimizing the relative contribution of leaky expression.\nComment 7:\nFigure 3a and page 14 – On page 14 the authors state that for the test molecule, there\nis “an EC50 of approximately 3 µM”. Looking at figure 3a, it looks like the EC50 should\nbe higher, closer to 10 or 11 µM. Should be double checked.\nResponse: We thank the reviewer for noting this error and apologize for the oversight.\nThe screenshot below shows the exact value as calculated by GraphPad Prism, where\nthe EC was determined to be 13.65 µM. This correction has been incorporated into the\nrevised manuscript on page 15, where the value is now accurately reported as 14 µM.\n₅₀\nEC50 shift, X is log(concentration)\nBest-fit values\nLogEC50Control 13.65 13.65\nEC50Ratio\nBottom 0.00279 0.00279\nTop 0.9729 0.9729\nHillSlope 0.1425 0.1425\nEC50Control 4.49225E+13 4.49225E+13\n95% CI (profile likelihood)\nLogEC50Control\nComment 8:\nFigure 3a – Since in CHAMP the signal is inversely proportional to the accumulation, it\nimportant to mention, in the text and/or figure legend, how the depicted plot was\nderived.\noversight. The legend for Fig. 3a has been revised to include details on how the plot was\nderived. It now reads:\n“The graph was obtained by plotting (1 – fold change, FC) against concentration, such\nthat a higher (1 – FC) value corresponds to greater molecular accumulation.”\nThis clarification has been incorporated into the revised figure legend in the manuscript.\nComment 9:\nFigure 5 – For the last panel, HBA (hydrogen bond acceptor) and HBD (hydrogen bond\nacceptor) should be defined in the legend for easy comprehension.\nResponse: We thank the reviewer for noting this omission and apologize for the\noversight. The legend for Fig. 5 has been updated to include definitions for the\nabbreviations HBA and HBD, which now read:\n“(HBA denotes hydrogen bond acceptor; HBD denotes hydrogen bond donor)\nThis clarification has been incorporated into the revised manuscript to ensure consistency\nand clarity in data presentation.\nComment 10:\nFigure 6 – Excellent analysis.\nResponse: We thank the reviewer for the feedback.\nComment 11:\nFigure 6i – In the legend of 6i it says “Permeability shift for compounds with or without\n(left) carbamate group, (middle) ester group, (right) carbamate”. There seems to be a\nmismatch between what is shown on the figure and what is described in the legend. The\nleft panel in the figure is labelled as “amide” not carbamate.\nfirst occurrence of “carbamate” in the legend of Fig. 6i has been corrected to “amide.”\nThis change is now reflected in the revised manuscript.\nComment 12:\nDiscussion, page 32 – “…are porin imbedded within the OM” should be “…is porins\nimbedded within the OM”.\nResponse: We thank the reviewer for noting this and apologize for the error. The\nsentence on page 33 has been corrected to read: “…is porins embedded within the outer\nmembrane (OM).” This correction has been implemented in the revised manuscript.\nComment 13:\nDefine replicates: In figures where experiments were carried out in replicates (such as\nfigures 2, 3, 4 and many supplementary figures), it is important to state if the\nreplicates were technical or biological in the legend of the corresponding figure.\nResponse: We thank the reviewer for noting this and apologize for the omission. In all\nfigures where experiments were performed in replicates, the phrase “of technical\nreplicates” has been added to the corresponding figure legends to clarify the nature of the\nreplicates used.\nReviewer #2\nComment 1:\nI found this study innovative and the experiments thoroughly take forward the concept.\nI thought that the data were strong and showed careful statistical analysis. There is one\nissue to address there: the microscopy in Fig.2 is not informative without showing\nalongside equivalent negative control images.\nbeen updated to include labeling of the cell wall using D-Lys(FITC)-OH (green), thereby\ndelineating the cell envelope (Angew Chem Int Ed Engl. 2012 Dec 7;51(50):12519-23.\ndoi: 10.1002/anie.201206749). Consequently, the cytosolic probe in this image was\nchanged to coumarin chloroalkane (Comcl), which emits in a distinct spectral window,\nallowing clear differentiation of the cytosol from the periplasmic space. In addition, E. coli\ncells that were not induced with IPTG but treated with Comcl have been included as a\nnegative control to establish the baseline fluorescence background.\nSimilarly, Fig. 2h and Fig. S4 have been updated to include E. coli cells that were not\ninduced with IPTG but treated with DBCOcl followed by coumarin azide (Comaz), to\nestablish the baseline fluorescence background. The use of Comaz was necessary\nbecause D-Lys(FITC)-OH (green) was employed to label the cell wall, thereby delineating\nthe cell envelope.\nFig. 2d. |\nFig. 2d and Fig. S1. | Confocal fluorescence images of E. coli cells expressing\nHaloTag, labeled with D-Lys(FITC) and treated with coumarin chloroalkane to\nproduce a blue cytosolic signal. E. coli cells harboring the plasmid pQE-30_HaloTag\n(Amp) were cultured overnight in LB medium. The overnight culture was diluted 1:100\ninto fresh LB medium supplemented with 500 µM D-Lys(FITC)-OH and incubated at 37\n°C with shaking until the optical density at 600 nm (OD ) reached approximately 1.2 to\nallow the incorporation of D-Lys(FITC)-OH in the cell wall. Protein expression was\n₆₀₀\ninduced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and cells were\nincubated for 1h to allow expression of the HaloTag protein. Uninduced cultures were\nmaintained as negative controls. Next, cells were washed three times with phosphate-\nbuffered saline (PBS) and incubated with 50 µM of coumarin chloroalkane (Comcl shown\non panel a) for 30 min at 37 °C with shaking. Cells were subsequently washed three times\nwith PBS, fixed with 4% (w/v) formaldehyde for 30 min, washed again and mounted on\n1% (w/v) agarose pads for imaging by confocal microscopy. Confocal fluorescence\nimaging was performed using a Leica STELLARIS 8 Super-Resolution Microscope\nequipped with a 100× oil-immersion objective. Image acquisition and analysis were\n11\ncarried out using ImageJ software. Excitation was performed using 405nm (Coumarin)\nand 488 nm (FITC) laser lines; panel b shows the images in different channels. Scale bar:\n2 µm.\nFig. 2h. |\nFig. S4. |\nFig. 2h. | Confocal fluorescence images of E. coli cells expressing HaloTag, labeled\nwith D-Lys(FITC) and treated with DBCOCl, which subsequently reacted with\nComaz to generate a blue cytosolic signal. E. coli cells harboring the plasmid pQE-\n30_HaloTag (Amp) were cultured overnight in LB medium. The overnight culture was\n12\ndiluted 1:100 into fresh LB medium supplemented with 500 µM D-Lys (FITC)-OH and\nincubated at 37 °C with shaking until the optical density at 600 nm (OD ) reached\napproximately 1.2 to allow the incorporation of D-Lys (FITC)-OH in the cell wall. Protein\n₆₀₀\nexpression was induced with 1 mM isopropyl β-D-1-thiogalactopyranoside (IPTG), and\ncells were incubated for 1h to allow expression of the HaloTag protein. Uninduced cultures\nwere maintained as negative controls. Next, cells were washed three times with\nphosphate-buffered saline (PBS) and incubated with 50 µM DBCOcl for 30 min at 37 °C\nwith shaking. Cells were subsequently washed three times with PBS, fixed with 4% (w/v)\nformaldehyde for 30 min, washed again, and incubated with 50 µM Comaz (on panel a)\nfor 1h at 37 °C with shaking. Finally, cells were washed three times and mounted on 1%\n(w/v) agarose pads for imaging by confocal microscopy Confocal fluorescence imaging\nwas performed using a Leica STELLARIS 8 Super-Resolution Microscope equipped with\na 100× oil-immersion objective. Image acquisition and analysis were carried out using\nImageJ software. Excitation was performed using 405nm (Coumarin) and 488 nm (FITC)\nlaser lines; panel b shows the images in different channels. Scale bar: 2 µm.\nComment 2 (Items 1 & 2):\nThe authors do a good job in considering the possible effect of the differential reactivity\nof azides with DBCO. The authors should comment in the Discussion on other\npossibilities for read-outs from the assay that do not precisely relate to the entry of the\nsmall molecule-azide of interest:\nPerhaps-\n1.the small molecule azide gets modified by the cell to some other compound\n-some molecule with different entry/exit properties\n-the azide could be reduced, so blocking reaction with DBCO\n2.the small molecule azide alters thiol levels in the cell which changes the non-specific\nareactivity of the cyclooctyne\nResponse: We thank the reviewer for this insightful comment. In response to queries 1\nand 2 above, additional sentences have been incorporated into the section on limitations\n13\n(page 31 of the revised manuscript). These additional sentences specifically address\npotential off-target effects and provide clarification regarding their possible impact on the\nstudy’s interpretations.\n“There is evidence that azides undergo modification within the biological milieu, a process\nshown to occur considerably more slowly for alkyl azides than for aryl azides, with\ncytosolic glutathione contributing to the reduction of azides to amines in E. coli. Although\nthe SPAAC DBCO–azide reaction is relatively slow, it proceeds several orders of\nmagnitude faster than the competing glutathione-mediated reduction of azides in\nbiological systems, making this competing process less disruptive to the overall\nfluorescence readout in our CHAMP assays.92-94 While it is likely that some small-\nmolecule azides undergo modification, we currently lack a method to quantify the extent\nof this effect. Nevertheless, since all azides in the panel are exposed to the same\nintracellular glutathione environment, the CHAMP assay provides a reliable measure of\nrelative accumulation across different azides under comparable conditions.”\nManuscript reference 92. Handlon AL, Oppenheimer NJ. Thiol reduction of 3'-\nazidothymidine to 3'-aminothymidine: kinetics and biomedical implications. Pharm Res.\nManuscript reference 93. Zhang C, Dai P, Vinogradov AA, Gates ZP, and Pentelute LB.\nSite-Selective Cysteine–Cyclooctyne Conjugation. Angew. Chem. Int. Ed. 2018, 57,\n6459.\nManuscript reference 94. Bertozzi CR, Sletten EM. Bioorthogonal Chemistry: Fishing\nfor Selectivity in a Sea of Functionality. Chem. Commun., 2013,49, 11007-11022\\\nComment 2 (Item 3):\nThe authors should comment in the Discussion on other possibilities for read-outs from\nthe assay that do not precisely relate to the entry of the small molecule-azide of interest:\nPerhaps-\n3. rapid lytic activity of the small molecule-azide\nexhibiting lytic activity may indeed undergo self-promoted uptake, resulting in relatively\nhigher intracellular accumulation. Conversely, certain compounds, such as polymyxin B\nnonapeptide (PMBN), primarily interact with the outer membrane—perturbing its structure\nand facilitating the uptake of other molecules without themselves entering the cytosol\n(Antimicrob. Agents Chemother. 38, 374–377 (1994)\nhttps://doi.org/10.1128/aac.38.2.374)\nWe acknowledge that some small-molecule azides could exhibit self-promoted uptake;\nhowever, this was not directly assessed in the present study. A commonly used method\nto evaluate this behavior is coincubation with a fluorescent small molecule, where\nenhanced uptake of the fluorophore indicates lytic activity. (Farmer, S., Li, Z. S. &\nHancock, R. E. J. Antimicrob. Chemother. 29, 27–33 (1992).\nOur CHAMP assay is responsive to molecules with lytic activity—a property we have\nleveraged to demonstrate enhanced uptake of Cy3az and Cy5az upon coincubation with\nPMBN (Fig. 2l). In future work, we aim to further exploit this capability of the CHAMP\nplatform to identify potential drug adjuvants—molecules that enhance uptake without\nnecessarily reaching the intracellular target themselves.\nComment 2 (Item 4):\nThe authors should comment in the Discussion on other possibilities for read-outs from\nthe assay that do not precisely relate to the entry of the small molecule-azide of interest:\nPerhaps-\n4. Engagement of the small molecule-azide with macromolecular targets hindering\nDBCO reaction.\nResponse: We agree with the reviewer’s observation. As noted by Bertozzi and\ncolleagues, it is indeed possible that some small-molecule azides undergo modification\nor interact with cellular components prior to reacting with DBCO. This represents an\ninherent limitation of our assay, and we currently lack a direct method to quantify or correct\nfor this effect. Nevertheless, since all azides are exposed to the same cellular\n15\nenvironment, the CHAMP assay provides a consistent and reliable measure of relative\naccumulation among azides, thereby allowing valid comparative analysis despite this\nconstraint. (Bertozzi, C. R. & Sletten, E. M. Bioorthogonal Chemistry: Fishing for\nSelectivity in a Sea of Functionality. Chem. Commun. 49, 11007–11022 (2013).\nComment 2 (Item 5):\nThe authors should comment in the Discussion on other possibilities for read-outs from\nthe assay that do not precisely relate to the entry of the small molecule-azide of interest:\nPerhaps-\n5.Potential impact of azide-bearing impurities in each small molecule-azide stock which\nmay have higher permeability.\nResponse: We appreciate the reviewer’s insightful comment. A paragraph addressing\ncompound purity as a limitation has been added to the manuscript on page 32. “The\ncommercial compounds used in this study are reported by the manufacturers to have a\npurity greater than 95%. However, we acknowledge that trace impurities within individual\nmolecules could influence the observed results. Given the large number of small\nmolecules screened in this work, we did not independently verify the purity of each\ncompound. To minimize any confounding effects from solvent interactions, the\nconcentration of DMSO—known to disrupt membrane integrity—was maintained below\n1% in all working solutions, in accordance with the guidelines of the Clinical and\nLaboratory Standards Institute and previous findings by Tunçer et al.” (Tunçer et al., Int.\nJ. Biol. Macromol., 2024, 267(Pt 2):131581. doi:10.1016/j.ijbiomac.2024.131581].\nComment 2 (Summary):\nThis is not a criticism of the approach but every assay has the potential for hits with\nalternative explanations, so it is good to contemplate such possibilities early on and\nconsider how to account for them.\nResponse: We thank the reviewer for the feedback.\nEven with a magnifying glass it is hard to see what is happening to the HaloTag in cells\nin Fig. 1d. It would be nice to edit the figure so this part is easier for readers to pick up.\nResponse: We thank the reviewer for this helpful observation. We agree that Fig.1d was\nnot sufficiently clear in its original form. The figures have now been revised and enhanced\nfor improved clarity and readability, ensuring that the data presentation is more intuitive\nto the reader.\nMinor point 2:\nAlso, it is not ideal that the fluorescence seems to be accumulating in Fig. 1D on the\noutside of the cell, which could lead to confusion.\nResponse: We appreciate the reviewer’s insightful observation that the previously\npresented image could give the impression that fluorescence was accumulating outside\nthe cell. To address this, the cartoons in Fig.1d (shown above) have been revised to more\naccurately represent intracellular localization. Specifically, dark green cytosolic\nfluorescence now denotes low-accumulating molecules. This change improves visual\nclarity and aligns the schematic with the experimental observations.\nMinor point 3:\nFig. 2l,m It would help readers to draw Cy5az and Cy3az in their most likely charge state\nResponse: We thank the reviewer for this helpful suggestion. Fig. 2l and Fig. 2m have\nbeen revised to include the most probable charge states of Cy5az and Cy3az expected\nunder the assay conditions. These updates improve the chemical accuracy of the\nillustrations and help readers better interpret the molecular behavior depicted.\nFig. 5c Give units for surface area\nResponse: Thank you for pointing this out, and we apologize for the earlier omission.\nThe y-axis in Fig. 5c has been updated to display the units for TPSA in square angstroms\n(Å²).\nMinor point 5:\nFig. 6g: Murcko and not Murko\nResponse: Thank you for bringing this to our attention, and we apologize for the\noversight. The x-axis label in Fig. 6g has been corrected to read “Murcko scaffolds” as\nshown below.\nMinor points 6:\nFig. 6i legend has carbamate twice\nResponse: We thank the reviewer for noting this and apologize for the confusion. The\nfirst occurrence of “carbamate” in the legend of Fig. 6i has been corrected to “amide.”\nThis change is now reflected in the revised manuscript.\nMinor points 7:\n“Interestingly, it was discerned that not all azides exhibited satisfactory reactivity within\nthe designated timespan of the assay, signifying that azide reactivity necessitates\ncareful consideration (Fig. S12). It is noteworthy that previous investigations have also\ndocumented variations in reactivity profiles between azides and strained\nalkynes,including DBCO.55”\nNice to have one or two sentences to summarize which kinds of azides react badly.\nadded a sentence on page 23 discussing azide reactivity that reads as follows: “Aromatic\nazides have been reported to exhibit significantly lower reactivity in strain-promoted\nazide–alkyne cycloaddition (SPAAC) reactions, with phenyl azide showing nearly a\nsevenfold reduction in reactivity compared to aliphatic azides, even when the aliphatic\nazide adopts an azidotoluene conformation. As for alkyl azides, the following trend in\nreactivity has been observed: primary > secondary > tertiary.” This statement is supported\nby prior studies (Dommerholt, J. et al. Highly accelerated inverse electron-demand\ncycloaddition of electron-deficient azides with aliphatic cyclooctynes. Nat. Commun. 5,\n5378 (2014). https://doi.org/10.1038/ncomms6378, and Zimmerman, E.S. et al.\nProduction of site-specific antibody–drug conjugates using optimized non-natural amino\nacids in a cell-free expression system. Bioconjug. Chem. 25(2), 351–361 (2014).\nReviewer #3\nComment 1 (Item 1):\nThe authors several times embellish the limitations of alternative methodologies and in\nsome cases the statements are plainly wrong. The authors' work should stand for itself\nwithout putting other approaches down and wrong facts should certainly not be used as\narguments for the superiority of their method. Some examples:\nPage 4/5: Subcellular localization can be assessed with MS, if proper cell fractionation\nprotocols with sufficient washing steps are followed. The same applies to fluorescent\ntags and inherent fluorescence, if used spectroscopically. Even more, the latter can be\nused in microscopy approaches, which, in most cases, easily allows assessment of\nsubcellular localization.\nResponse: We thank the reviewer for this thoughtful observation and apologize for the\nuse of overly strong language in describing the challenges associated with assigning\nmolecular localization in E. coli. The text has been revised for clarity and tone, and now\nreads on page 5:\n19\n“(b) would ambiguously define molecule localization unless additional, careful\nfractionation methods are included to show periplasmic or cytoplasmic accumulation.”\nThis revision more accurately reflects the methodological nuances involved without\noverstating the limitation.\nComment 1 (Item 2):\nPage 9: There simply is no challenge to distinguish membrane from cytosol in bacteria.\nThe difference between cell surface association, membrane localization, and cytosolic\nlocalization are glaringly obvious, even with simple wide-field microscopes. Even the\nouter and inner membrane can be easily distinguished from each other when following\nappropriate protocols. The authors even show this themselves with abundantly clear\ncytosolic fluorescence of their probe.\nPlease check through the manuscript and remove faulty claims.\nResponse: We appreciate the reviewer’s insightful comment. In response, we have\nrevised the text to remove language referring to the difficulties in distinguishing cytosolic\nand periplasmic localization and have replaced it with a clear explanation of how the\ncytosol was delineated. The revised text now reads on page 9:\n“To confirm that the accumulation measurements in this study reflect cytoplasmic\nlocalization of molecules, the cytoplasmic compartment of E. coli cells was delineated by\nlabeling the cell wall with D-Lys(FITC)-OH, which emits green fluorescence (Angew Chem\nInt Ed Engl. 2012 Dec 7;51(50):12519-23. doi: 10.1002/anie.201206749). Consequently,\nthe cytosolic probe for confocal imaging was changed to coumarin chloroalkane (Comcl),\nwhich fluoresces in a distinct spectral window, thereby enabling clear spatial separation\nof cytosolic and periplasmic signals. As a control, E. coli cells that were not induced with\nIPTG but treated with Comcl were included to establish the baseline fluorescence\nbackground.\nThis revision provides a more direct and mechanistic explanation of how cytoplasmic\nlocalization was established in the assay.\nThe authors describe the SYTOX green assay wrong (page 11). Firstly, SYTOX dyes\nare perfectly cell wall permeable. The cell wall is the peptidoglycan layer, not the whole\ncell envelope structure built up of inner membrane, peptidoglycan cell wall, and outer\nmembrane. In order to penetrate, SYTOX dyes need both the outer and inner\nmembrane compromised. Thus, they indicate disruption of membrane (not cell wall)\nintegrity. Further, since the inner membrane needs large pores to allow the passage of\nthe dye, which will kill the cells, SYTOX dyes indicate dead cells. So what the authors\nhave proven here is simply that Halo tag expression does not simultaneously create\nlarge holes in the inner and outer membrane of E. coli. Please rephrase this part to\ncorrect this.\nResponse: Thank you so much for pointing this out and we are sorry for the\nmisrepresentation of the SYTOX Green dye. The SYTOX Green paragraph now appears\non page 12 and has been modified to read: “To verify the viability of the E. coli cells after\ninducing HaloTag expression and subjecting cells to DBCOcl treatment, we conducted a\nSYTOX Green analysis.46 In live prokaryotic cells, SYTOX Green is excluded from cells\nwith intact permeability barriers.47 Our results revealed that the expression of HaloTag\nand incubation of these cells with DBCOcl and its occupancy by HaloTag in E. coli did not\ncompromise the integrity of the bacterial cytoplasmic membrane (Fig. S4). This finding\nemphasizes the suitability of utilizing CHAMP for investigating molecular permeation to\nthe cytosol without causing disruption to the cell’s permeability barriers.”\nComment 2 (Item 2):\nA second issue with this assay is that it simply does not support the authors'\nconclusions. Because SYTOX dyes need both the inner and outer membrane\ncompromised, a negative result cannot exclude that either one of them may be\ndamaged. If the authors truly want to make the claim that the cell envelope as a whole\nis not damaged, they must assess inner and outer membrane integrity separately. This\ncould, for example, be achieved using NPN for the outer and DiSC( )5 for the inner\n3\n21\nmembrane.\nResponse: We thank the reviewer for this valuable suggestion. The manuscript has been\nrevised accordingly to present conclusions derived solely from the SYTOX Green assay.\nAdditionally, in the Supplementary Information (page S8), the term “integrity” has been\nreplaced with “viability”, as reflected in the updated caption: “Fig. S4. Cellular viability\nexamination”. We greatly appreciate the reviewer’s recommendation regarding the NPN\nand DiSC( )5 dyes as a tool and we plan to incorporate these assays in future studies to\n3\nevaluate changes in outer and inner membrane integrity.\nComment 3:\n3. The authors misrepresent/misinterpret vancomycin. On page 18, they say that no\nintracellular accumulation of vancomycin was observed, which would be consistent with\nthe lack of biological activity against E. coli. Vancomycin binds to lipid II, precisely to the\nD-Ala-D-Ala motif of the pentapeptide chain. This is a cell surface structure that is\npresented on the outer leaflet of the inner membrane. Naturally, vancomycin would\nnever be found intracellularly, not even in sensitive bacteria. However, vancomycin is\nperfectly able to bind to lipid II and inhibit E. coli, if the outer membrane is compromised.\nThe authors even show this themselves (page 19). The authors further imply (by\nlumping vancomycin in with puromycin and others) that what they observe is uptake into\nthe intracellular space. As outlined above, that is not the case with vancomycin, which\nbinds to the inner membrane.\nPlease correct these errors and also generally pay more attention to distinguishing cell\nand cell envelope structures and defining uptake appropriately (as in uptake into the\nperiplasm or cytoplasm) as this is of crucial relevance to the content of the paper.\nResponse: We thank the reviewer for pointing this out and apologize for the lack of clarity\nin our discussion of vancomycin. Our laboratory has studied and published extensively\non vancomycin and its effects on cell wall biosynthesis, but in attempting to provide a\nbrief\nremark on vancomycin accumulation, the phrasing inadvertently came across as\n22\ninaccurate (https://doi.org/10.1002/anie.201704851 and\nhttps://doi.org/10.1039/c7sc02721d). The text has been revised to more accurately reflect\nthe established mechanism of vancomycin activity and its lack of accumulation in E. coli\ndue to the outer membrane barrier.\nThe text has now been edited to read on page 19:\n“Of note, there was no detectable accumulation of vancomycin azide (vanAZ1) within the\ncytosolic space of E. coli, consistent with expectations for this organism. Vancomycin\nexerts its activity by binding to the D-Ala-D-Ala terminus of the lipid II peptidoglycan\nprecursor, which is presented on the outer leaflet of the inner membrane (Molinari, H. et\nal. Structure of vancomycin and a vancomycin/D-Ala-D-Ala complex in solution.\nBiochemistry 1990, 29, 9, 2271–2277 https://doi.org/10.1021/bi00461a010) For\nvancomycin to accumulate in the cytoplasm of E. coli, it would necessarily have to\ntraverse the periplasmic space, where it would encounter its target, thereby rendering the\ncells sensitive to the antibiotic. However, E. coli—and, more broadly, Gram-negative\nbacteria—are intrinsically resistant to vancomycin due to the impermeability of their outer\nmembrane and only become susceptible when this barrier is disrupted (Vaara, M. et al.\nSusceptibility of gram-negative bacteria to polymyxin B nonapeptide. Antimicrobial\nAgents and Chemotherapy, 01 Jun 1984, 25(6):701-705.\nhttps://doi.org/10.1128/aac.25.6.701)\nAccordingly, vancomycin serves as an effective negative control in our CHAMP assay for\nassessing the intracellular accumulation of other antibiotics.”\nComment 4:\nIn connection with that, the authors (wrongly) remark that microscopy would pose\ndifficulties in distinguishing the membrane from the cytosol, but their method clearly\ncannot achieve this as outlined above. I would be really curious to see microscopy\nassays here. For example, vancomycin and puromycin uptake in WT compared to ∆tolC\nwould be a good setup to show (i) whether their method os also be well-suitable for\nmicroscopy (which would be really exciting in my opinion) and (ii) whether it actually can\ndistinguish subcellular localization.\nperformed such an investigation, we believe that the CHAMP assay would be compatible\nwith confocal microscopy and could be used to compare fluorescence accumulation in\ncells treated with a molecule such as puromycin in both WT E. coli and ΔtolC E. coli. In\nsuch a setup, it is expected that higher intracellular accumulation would yield a dimmer\nfluorescence signal.\nWhile confocal microscopy provides excellent spatial resolution, it is limited in throughput\nand quantitative sensitivity when detecting subtle differences between molecules with\ncomparable, though not identical, accumulation profiles. For this reason, our CHAMP\nassays utilize flow cytometry, which enables high-throughput analysis of approximately\n10,000 cells per sample. This approach allows for the simultaneous evaluation of up to\n10 different conditions, providing reliable relative accumulation measurements in under\nfive minutes.\nComment 5:\nSince the Halo tag is expressed in the cytosol, I am wondering how it could detect\nvancomycin uptake (which sits in the membrane as outlined above). I can imagine four\nscenarios: (i) vancomycin flips over the membrane to the inner leaflet and is then able to\ninteract, (ii) the Halo tag is somehow also expressed in the periplasm where it would\nencounter vancomycin, (iii) the azide label alters vancomycin properties enabling\ncytosolic translocation, or (iv) the experimental conditions do compromise the inner\nmembrane so vancomycin and/or Halo tag can cross this barrier. Each one of these\nwould have rather crucial implications for the usability of the method and I would\nstrongly suggest to address this.\nResponse: We appreciate the reviewer’s observation regarding the inclusion of\nvancomycin in our assays. For molecules that do not penetrate the cytosol, such as\nvancomycin, the CHAMP assay does not provide sufficient information to determine\nsubcellular localization. Vancomycin was included in our study as a negative control to\nillustrate this limitation.\nthe cytosol. Consequently, only molecules that reach the cytosol can be detected in the\nCHAMP assay, as their presence is reflected by a decrease in fluorescence intensity\nrelative to the untreated control. Conversely, if a molecule yields a fluorescence signal\ncomparable to that of the untreated control, it may have localized to the periplasm or\nremained outside the cell; however, the assay cannot distinguish between these\nscenarios. Thus, the CHAMP assay specifically reports cytosolic accumulation but\nremains agnostic to molecular localization outside the cytosol.\nComment 6:\nSomewhat related to the same question, I would be really curious if periplasmic\nlocalization would be possible to see. Would the system work with periplasmic\nexpression?\nResponse: We thank the reviewer for this thought-provoking question. The CHAMP\nassay can indeed be adapted for use with HaloTag expressed in the periplasm by\nemploying appropriate localization signal sequences to direct protein targeting. While this\nline of investigation is currently ongoing in our laboratory, it was not included in the present\nstudy. We anticipate presenting these findings in future publications.\nComment 7:\nThe authors make some mistakes describing PMBN. Firstly, they keep talking about\nsublethal concentrations (page 12/13), which implies that PMBN is lethal. However, it\nhas no activity against E. coli (under standard laboratory conditions) that would show in\nstandardized MIC assays (MICs usually above 512 µg/mL). Secondly, they state that it\ncould interfere with the inner membrane (page 20). However, that is not the case (see\nfor example PMID 36165741). The reference they cite for this does not state this either.\nPlease correct this.\nlethal” has been corrected to “low.” Indeed, PMBN does not disrupt the inner membrane\nat the 10 mM concentration used in our assays. We have confirmed this experimentally\nusing the membrane potential-sensitive dye DiSC( )5 as shown below.\nE. coli cells induced to express HaloTag protein were treated with 10 µM PMBN or 0.2%\n₅\nTriton X-100 for 30 min; untreated cells served as a negative control. A decrease in\nDiSC( )5 fluorescence in PMBN-treated cells indicates quenching due to dye\n3\nintercalation into the membrane, whereas dequenching occurs upon pore formation and\nloss of membrane potential as the dye redistributes into the cytoplasm. Cellular\nfluorescence was measured using a plate reader (excitation = 622 nm; emission = 670\nnm). Data are presented as mean ± s.d. (n = 3 independent samples within a single\nexperiment). Statistical significance was determined using a two-tailed t-test (P < 0.05,\n*P < 0.01, **P < 0.001, ***P < 0.0001; ns = not significant).\nComment 8:\nAlso regarding this, on page 20 where the authors argue with the possible inner\nmembrane effects of PMBN, it would make much more sense to argue that FhuA has a\nlower substrate range because it is a porin, which is naturally limited in the\nsize and charge of molecules that it lets pass. PMBN induces large-scale destruction of the OM,\nletting much larger molecules through including massive molecules like nisin.\ntypically transports substrates in the range of 700–1000 Da, including ferricrocin,\nferrichrome, albomycin, and rifamycin CGP 4832 (Ferguson, A. D. et al. Science 282,\n2215–2220 (1998); https://doi.org/10.1126/science.282.5397.2215). In this study, we\nutilized an open, non-selective variant of FhuA—designated “pore”—which does not\ndiscriminate based on substrate hydrophilicity and is localized to the outer membrane.\nAlthough the exact substrate size range for this variant has not been fully characterized,\nE. coli K-12 BW25114 expressing the pore form of FhuA has previously been shown to\ndisplay a 16-fold reduction in the minimum inhibitory concentration (MIC) of vancomycin\ncompared to the wild-type strain, indicating increased permeability (Krishnamoorthy, G.\net al. Antimicrob. Agents Chemother. 60, 7372–7381 (2016);\nhttps://doi.org/10.1128/AAC.01882-16).\nOn page 21, the Main Text has been revised to read “because it is a porin. In this work,\nan open and non-selective variant of FhuA that does not discriminate based on\nhydrophilicity, designated “pore” was used; the pore is localized to the outer membrane.\nThe substrate size range for this strain is not clear but E. coli K12 BW25114 expressing\nthis variant of FhuA was shown to be sensitive to vancomycin with a 16-fold reduction in\nMIC relative to the wild type strain”.\nComment 9:\nOn page 28 the authors argue with using MIC changes as proxy for uptake. While the\nargument that MICs depend on target interaction is of course true, it is also weak in the\ncontext of the paper as the authors argue that structural changes to compounds will\naffect target interaction. However, that is also the case for the azide linkers used here.\nThe argument could further be broken down to testing the activity of compounds with\nand without PMBN or in WT and a permeable mutant, which is the same setup the\nauthors are using here and hence not really that different as a concept.\nResponse: We thank the reviewer for this insightful comment. We agree that a minimum\ninhibitory concentration (MIC) assay is a valuable method for assessing the arrival of\n27\nmolecules at their intracellular drug target sites. For our azido-tagged molecules,\nconducting MIC measurements in the presence or absence of PMBN, or using WT and\nFhuA mutant strains, provides an effective way to account for structural modifications in\nthe small-molecule drugs under investigation. In light of this, we have revised the\nparagraph to read on page 29: “CHAMP adds a tool to the existing methods used to\nmeasure accumulation of molecules in the cytosol.”\nComment 10:\nSimilarly, on page 30/31 the authors argue for the usability of their method in screening\nfor potentiators that disrupt outer membrane integrity. While I do not doubt that this\ncould an application, the argument is also weakened by the observation that their assay\nis itself impacted by the integrity of the outer membrane, making such measurements\nhard to calibrate.\nResponse: We thank the reviewer for pointing out this apparent contradiction. In the\nCHAMP assay, potentiators of molecular accumulation can indeed be identified using\nazido-tagged dyes that exhibit low fluorescence signal-to-noise ratios in the absence of a\nmembrane-disrupting agent. A representative example is shown in Fig. 2l, where sulfo-\nCyanine3 azide and sulfo-Cyanine5 azide do not accumulate in E. coli above background\nlevels when incubated with vehicle alone. However, in the presence of a membrane\ndisruptor such as PMBN, a marked increase in fluorescence intensity is observed,\nindicating enhanced intracellular accumulation. It follows, therefore, that experimental\nconditions producing a fluorescence profile similar to that observed with PMBN—when\nsulfo-Cyanine dyes are co-incubated with a test molecule—would suggest that the\nmolecule possesses membrane-disrupting or permeability-enhancing activity.\nComment 11:\nRelated to the last point, I would wish that the authors would more honestly and critically\nreflect on the limitations of their method and in an open and honest way compare the\npros and cons between their method and existing ones. The embellishment of the\nlimitations of competing techniques and the absence of honest reflections on their\n28\nmethod’s limitations is a red thread in the paper that I cannot condone.\nResponse: We thank the reviewer for a thorough review of our work, and we have\nedited the original manuscript to highlight limitations inherent in the assay as follows:\nPage 20 “ Several antibiotics act within the periplasmic space of E. coli and other\nGram-negative bacteria, including β-lactam antibiotics, which target periplasmic\npenicillin-binding proteins (PBPs) involved in peptidoglycan synthesis, and\nglycopeptide antibiotics such as vancomycin, which inhibit cell wall cross-linking.66-\nlocalization represents a limitation of the assay in its present form.”\nPage 23 “Aromatic azides have been found to have lower reactivity in SPAAC\nreactions with nearly a sevenfold reduced reactivity of phenyl azide in comparison\nwith aliphatic azides even where the aliphatic azide conformation is in the form of\nazidotoluene. As for alkyl azides, the following trend in reactivity has been\nobserved: primary > secondary > tertiary. This statement is supported by prior\nstudies.72 It was quickly recognized that variations in the intrinsic reactivities of\nazides would constrain the analysis to only those azides exhibiting comparable\nreactivity, thereby narrowing the chemical diversity and reducing the overall\nscreening breadth achievable with the CHAMP platform.”\nPage 31-32 “We acknowledge that there are limitations to the CHAMP assay.\nPrincipally, in this iteration, the target pathogen must be amenable to genetic\nmanipulation, and the molecule must be tagged with an azide. There is evidence\nthat azides undergo modification within the biological milieu, a process shown to\noccur considerably more slowly for alkyl azides than for aryl azides, with cytosolic\nglutathione contributing to the reduction of azides to amines in E. coli. Although the\nSPAAC DBCO–azide reaction is relatively slow, it proceeds several orders of\nmagnitude faster than the competing glutathione-mediated reduction of azides in\nbiological systems, making this competing process less disruptive to the overall\nfluorescence readout in our CHAMP assays.92,93,94 ”\nsmall molecule-azide library stock which may skew permeability of the molecules\nunder test. Given the extensive library of small molecules screened in this work,\nwe did not independently assess the purity of each compound.”\nReviewer #4\nComment 1:\n- The manuscript is difficult to read for outsiders. First, I suggest a more basic scheme\nabout the function bringing the subject to a broader audience.\nResponse: We thank the reviewer for this helpful suggestion to simplify the presentation\nof the manuscript. Accordingly, we have restructured the cartoons, graphed data, figure\ncaptions, confocal microscopy images, and corresponding main text descriptions to\nimprove clarity and ensure that the information is presented in a more intuitive and reader-\nfriendly manner.\nComment 2:\n- What is the sensitivity, how many bacteria are needed? Could this extend to single\ncells?\nResponse: We appreciate the reviewer’s comment regarding single-cell analysis. The\nprimary challenge with applying this method to individual cells lies in the practical difficulty\nof subjecting a single cell to multiple incubation, centrifugation, and washing steps while\nreliably recovering that same cell for laser interrogation in the flow cytometer. Given that\ncells are inevitably lost during these preparative steps, we employ flow cytometry analysis\nof approximately 10,000 events as a representative population to ensure statistical\nrobustness when comparing treatments.\nThe accompanying graph illustrates that even when smaller subsets of cells are analyzed,\nthe relative differences between HaloTag-expressing cells (IPTG-induced) and non-\nnon-\ninduced controls remain consistent. In this experiment, cells induced with 1 mM IPTG to\n30\nexpress HaloTag protein were sequentially labeled with DBCOcl (30 min) and R110az (50\nµM, 1 h), followed by flow cytometric analysis using varying event count thresholds. The\ndata represent technical replicates from six independent experiments, with uninduced\ncells serving as negative controls.\nAnalysis of mean fluorescence intensity (MFI) using one-way ANOVA, with the 10,000-\nevent dataset as the control, revealed no statistically significant differences between\ndatasets collected from smaller sample sizes. However, examination of fluorescence\nintensity distributions using violin plots indicated substantial variability among individual\nevents, including outliers with extremely high or low signals. Such outliers, if\ndisproportionately represented in smaller datasets, could lead to misleadingly high or low\nfluorescence readings. For this reason, we consider smaller event counts to be unreliable\nfor quantitative comparisons and therefore rely on 10,000-event analyses for consistent\nand representative measurements.\nTo determine the minimum number of cells required to reliably detect differences in\nmolecular accumulation, E. coli cells induced to express the HaloTag protein were treated\nsequentially with DBCO–Cl followed by 50 µM R110–azide for 1 h. Uninduced cells\nserved as negative controls. Cellular fluorescence was quantified using flow cytometry\nwith excitation at 488 nm and emission at 525 nm. Flow cytometer settings were adjusted\nto record 10,000, 5,000, 1,000, 500, 100, 10, or 1 events per sample. Fluorescence\nintensity data were analyzed as the mean of six technical replicates for each stopping\ncondition. Results are presented as mean ± SD from three independent experiments.\nStatistical analysis was performed using a one-way ANOVA, with the mean fluorescence\nvalue from 10,000 events serving as the reference group.\nNo statistically significant differences were observed across the event groups (P = 0.767),\nindicating that acquisition of fluorescence data from as few as 500 cells provides a reliable\nestimate of accumulation comparable to that obtained from 10,000 events. Statistical\nsignificance was defined as (*P < 0.05, **P < 0.01, ***P < 0.001, ***P < 0.0001, ns = not\nsignificant).\nTo determine the minimum number of cells required to reliably capture differences in\nmolecular accumulation, E. coli cells induced to express the HaloTag protein were treated\nwith DBCO–Cl, followed by 50 µM R110–azide for 1 h. Uninduced cells served as\nnegative controls. Cellular fluorescence was quantified by flow cytometry using excitation\nat 488 nm and emission at 525 nm. The flow cytometer was configured to record 10,000,\n5,000, 1,000, 500, 100, 10, or 1 event(s) per acquisition. The resulting fluorescence\ndistributions are presented as a violin plot, with each event represented along the y-axis\naccording to fluorescence intensity. Data are expressed as mean ± SD from three\nindependent experiments. Statistical significance was assessed by one-way ANOVA,\nusing the 10,000-event dataset as the reference group. No significant differences were\nobserved across event counts (P = 0.767), indicating that fluorescence data derived from\nas few as 500 cells are sufficient to convey reliable information on accumulation\ndifferences. Statistical significance was defined as (*P < 0.05, **P < 0.01, ***P < 0.001,\n***P < 0.0001, ns = not significant).\nComment 3:\n- I congratulate the authors for the amount of work and test. This is a very good basis\nfor other works. However, I read the ms several times to catch the bigger picture.\nMaybe restructuring might render the ms easier to read? (I understand that its not\nthe primary goal to make it easy).\nResponse: We thank the reviewer for this helpful suggestion to simplify the presentation\nof the manuscript. Accordingly, we have restructured the cartoons, graphed data, figure\ncaptions, confocal microscopy images, and corresponding main text descriptions to\nimprove clarity and ensure that the information is presented in a more intuitive and reader-\nfriendly manner.\nComment 4:\nPlease clarify the limits of the approach.\nrevised the Main Text to explicitly address additional limitations of the CHAMP assay.\nAmong others, we now highlight how differences in azide–DBCO reactivity can affect\nassay reliability, particularly when comparing the intracellular accumulation of chemically\ndistinct azides. Although this limitation is partially mitigated by testing each molecule in\ntwo biological contexts, variability in chemical reactivity remains an important factor to\nconsider.\nTo clarify these points, we have added the following text in the manuscript as shown\nbelow:\nPage 20 “ Several antibiotics act within the periplasmic space of E. coli and other\nGram-negative bacteria, including β-lactam antibiotics, which target periplasmic\npenicillin-binding proteins (PBPs) involved in peptidoglycan synthesis, and\nglycopeptide antibiotics such as vancomycin, which inhibit cell wall cross-linking.66-\nlocalization represents a limitation of the assay in its present form.”\nPage 23 “Aromatic azides have been found to have lower reactivity in SPAAC\nreactions with nearly a sevenfold reduced reactivity of phenyl azide in comparison\nwith aliphatic azides even where the aliphatic azide conformation is in the form of\nazidotoluene. As for alkyl azides, the following trend in reactivity has been\nobserved: primary > secondary > tertiary. This statement is supported by prior\nstudies.72 It was quickly recognized that variations in the intrinsic reactivities of\nazides would constrain the analysis to only those azides exhibiting comparable\nreactivity, thereby narrowing the chemical diversity and reducing the overall\nscreening breadth achievable with the CHAMP platform.”\nPage 31-32 “We acknowledge that there are limitations to the CHAMP assay.\nPrincipally, in this iteration, the target pathogen must be amenable to genetic\nmanipulation, and the molecule must be tagged with an azide. There is evidence\nthat azides undergo modification within the biological milieu, a process shown to\n34\noccur considerably more slowly for alkyl azides than for aryl azides, with cytosolic\nglutathione contributing to the reduction of azides to amines in E. coli. Although the\nSPAAC DBCO–azide reaction is relatively slow, it proceeds several orders of\nmagnitude faster than the competing glutathione-mediated reduction of azides in\nbiological systems, making this competing process less disruptive to the overall\nfluorescence readout in our CHAMP assays.92,93,94 ”\nPage 32 “An additional consideration is the potential impact of impurities in each\nsmall molecule-azide library stock which may skew permeability of the molecules\nunder test. Given the extensive library of small molecules screened in this work,\nwe did not independently assess the purity of each compound.”\nWe thank the reviewers for their careful evaluation of this manuscript and attention to\ndetail. We feel that the suggested revisions have helped us craft an improved manuscript\nthat will better serve the community.\nSincerely,\nMarcos Pires\nProfessor of Chemistry\nDepartment of Chemistry\nUniversity of Virginia\nCharlottesville, VA 22904\nEmail: mpires@virginia.edu\n35"}, {"round": 2, "reviewer_comments": {"reviewer_1": "My concerns and comments have been addressed sufficiently by the authors in the revisions process and I recommend this\nmanuscript for publication.", "reviewer_2": "All my comments have been well answered and I recommend accepting the paper.", "reviewer_3": "The authors have written a very careful and well-structured rebuttal, amended or corrected the manuscript where necessary,\nand brought forward good arguments where reviewer suggestions were not or only partly followed. For my comments, I am\nsatisfied with the authors' edits and replies and I have no further revisions. I recommend acceptance of the manuscript.", "reviewer_4": "The author replied to the question. In my opinion their approach is highly interesting for the field and I recommend\npublication as it is."}, "author_response": ""}]

Decision letter