Capillary zone electrophoresis with fraction collection for separation

Subscriber access provided by University of Sussex Library

Article

Capillary zone electrophoresis with fraction collection for separation, culturing, and identification of bacteria from an environmental microbiome Bonnie Huge, Matthew M. Champion, and Norman J. Dovichi Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/acs.analchem.8b05984 • Publication Date (Web): 25 Feb 2019 Downloaded from http://pubs.acs.org on March 6, 2019

Just Accepted “Just Accepted” manuscripts have been peer-reviewed and accepted for publication. They are posted online prior to technical editing, formatting for publication and author proofing. The American Chemical Society provides “Just Accepted” as a service to the research community to expedite the dissemination of scientific material as soon as possible after acceptance. “Just Accepted” manuscripts appear in full in PDF format accompanied by an HTML abstract. “Just Accepted” manuscripts have been fully peer reviewed, but should not be considered the official version of record. They are citable by the Digital Object Identifier (DOI®). “Just Accepted” is an optional service offered to authors. Therefore, the “Just Accepted” Web site may not include all articles that will be published in the journal. After a manuscript is technically edited and formatted, it will be removed from the “Just Accepted” Web site and published as an ASAP article. Note that technical editing may introduce minor changes to the manuscript text and/or graphics which could affect content, and all legal disclaimers and ethical guidelines that apply to the journal pertain. ACS cannot be held responsible for errors or consequences arising from the use of information contained in these “Just Accepted” manuscripts.

is published by the American Chemical Society. 1155 Sixteenth Street N.W., Washington, DC 20036 Published by American Chemical Society. Copyright © American Chemical Society. However, no copyright claim is made to original U.S. Government works, or works produced by employees of any Commonwealth realm Crown government in the course of their duties.

Page 1 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Analytical Chemistry

Capillary zone electrophoresis with fraction collection for separation, culturing, and identification of bacteria from an environmental microbiome Bonnie Jaskowski Huge 1, Matthew M. Champion 1,2, Norman J. Dovichi 1,2,*

1Department

of Chemistry and Biochemistry, and 2Advanced Diagnostics and Therapeutics, University of Notre Dame, Notre Dame, IN 46556 USA *email: [email protected]

ACS Paragon Plus Environment

Analytical Chemistry 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

ABSTRACT: Capillary zone electrophoresis (CZE) can produce high-resolution separations of biological samples, including microbial mixtures. The study of complex populations of microorganisms using CZE is limited because most detectors have limited sensitivity, are destructive, and provide limited information for microbial identification. To address these issues, we developed an integrated capillary zone electrophoresis apparatus to fractionate bacteria from complex mixtures. We deposited fractions onto nutrient agar in a Petri dish for microbial culturing, and we subjected the observed colonies to Sanger sequencing of a phylogenetic marker, the 16S rRNA gene, for microbial identification. We separated and cultured both a single bacteria species, the model Gram (-) organism Escherichia coli, and a complex environmental isolate of primary sewage effluent. Sequence analysis of the 16S rRNA genes from this mixture identified 15 ± 5 distinct bacterial species per run. This approach requires minimal manipulation of microbial populations and combines electrophoretic fractionation of bacterial cells with automated collection for accurate identification of species. This approach should be applicable to microorganisms in general, and may enable discrimination of physiologically different cells in complex assemblages, such as in microbiome samples.

Page - 1


Page 2 of 26

Page 3 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


An environmental microbiome is a complex community that contains hundreds or thousands of microbial species and associated viruses. Characterization of a microbiome is challenging because only a small fraction of the species in the community is readily cultured, so that most microbial species are invisible to classic culture-based microbiological methods (1-2). Various physicochemical and molecular measurements are employed to characterize these microbes and microbial communities. Among these methods, the behavior of bacteria in an electric field and during electrophoresis was reported in the early 20th century (3-11), and instrumentation was commercialized for measurement of the electrophoretic mobility and zeta potential of single microbes (12). This early instrumentation tended to be cumbersome, and a small community of researchers investigated the use of capillary zone electrophoresis (CZE) to simplify microbial characterization. Ebersole and McCormick performed a pioneering study that demonstrated the use of CZE for the baseline separation of two pure bacterial cultures (13). Fractions were manually collected after separation, were of high purity, and retained viability. Armstrong and others followed this work with publications that reported the characterization of more complex bacterial mixtures using CZE (14-25). Under certain combinations of background electrolyte and bacterial loading amounts, these separations demonstrated remarkably sharp and well-resolved peaks. Several of these studies considered the electrophoretic behavior of pathogens (16, 18, 21-23). Others were developed as general detectors of microbial contamination; these studies took advantage of electrophoretic conditions that induced microbial aggregation (14, 17, 19, 21, 25). These studies suffered from three limitations. First, manual manipulations were required to collect fractions. Second, simple optical detectors, such as UV absorbance or laser-induced fluorescence, were used to detect the separated bacteria, which provide very little specificity.

Page - 2



However, in one notable case, capillary electrophoresis was combined with in situ hybridization to specifically detect a target species in a binary mixture (18). Third, bacterial aggregation was frequently observed at high sample concentrations. This aggregation destroys resolution of species, but, as noted above, can be used to screen samples for the presence of any bacterial contamination, albeit without identification of the contaminant. In this paper, we report the first coupling of capillary zone electrophoresis with an automated fraction collector for separation of bacteria from a complex microbiome, the deposition of those fractions onto a Petri dish for culturing, and the Sanger sequencing of the 16S gene from the culturable organisms in that microbiome for detection. Capillary electrophoresis provides separation of the complex microbiome into fractions to simplify subsequent molecular analysis, culturing of fractions provides powerful amplification of the separated microbes, and the sequencing of a phylogenic marker gene provides characterization of these culturable microbial species.

Page - 3


Page 4 of 26

Page 5 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


EXPERIMENTAL SECTION Materials and Methods Except as noted, all reagents were analytical grade and purchased from Sigma-Aldrich. Solutions were prepared from deionized-distilled water (ddH2O) obtained from a Barnstead Nanopure System (Thermo-Fisher Scientific). Culture-related consumables and glassware were purchased sterile (Corning and VWR) or autoclaved prior to use. Model Strain and Bacterial Culture  E. coli HB101 obtained from Coli Genetic Stock Center (CGSC at Yale University) was used to develop our separations. The strain was chemically-transformed with a GFP-expressing plasmid (pglo, Bio-Rad), under control of the pBAD arabinose promoter to aid in visualization of colonies. Bacteria were cultured using LB medium (Miller’s LB powder) supplemented with 100 μg/mL ampicillin in culture tubes at 37 °C at 100 rpm overnight. Fresh LB medium supplemented with 100 μg/mL ampicillin was inoculated with the overnight cultures (1:100 dilution) in shaking flasks and incubated at 37 °C at 100 rpm until they reached mid-exponential growth; 4 h total, 0.2% L-arabinose (Teknova) (v/v) was added to liquid cultures at 2 h. After growth (approx. ~2h), cultures were harvested by centrifugation (5000 x g 10 min) and washed three times (1x culture volume) with sterile-filtered PBS (Dulbecco’s Phosphate Buffered Saline). Washed cells were re-suspended in sterile PBS for subsequent analysis. Wastewater Sampling and Long-term Storage  A 2 L aliquot of primary effluent was collected at the Mishawaka Wastewater Treatment Facility (Mishawaka, Indiana, USA). The sampling site was post-settling and pre-chlorination. Microorganisms were isolated by centrifugation (5000 x g 10 min) and washed with sterile-

Page - 4



filtered PBS: the wastewater was centrifuged in ~40 mL aliquots in 50 mL centrifuge tubes. The pelleted material was combined in the washing steps until all pellets were combined into one tube. The final pellet was suspended in PBS with glycerol (22%) for long-term storage at -80° C. Aliquots were thawed and diluted in sterile PBS prior to analysis. Instrumentation The electrophoresis system was equipped with a 60 cm long bare fused capillary (100 μm ID, 160 μm OD, Polymicro Technologies) that was inserted into an injection block, similar to a published design (26), supplied with high voltage for electrophoresis (Spellman CZE1000R). We constructed an instrument to interface capillary zone electrophoresis with culturebased detection using automated fraction collection. The basic instrument is described in detail elsewhere (27-28). Briefly, the distal end of the separation capillary was threaded through a Tee fitting and positioned at the exit of a precision dispensing nozzle (The Lee Company). For automated fraction collection, the nozzle was secured above a motorized microscope stage (Prior Scientific), which holds a collection plate or Petri dish. The instrument was controlled with software written in Labview (See supporting information). Electrophoretic Fractionation  The separation capillary was conditioned and flushed with methanol (MeOH), ddH2O, 1 M NaOH, ddH2O, and 10 mM Tris-HCl (pH 7.5) in series prior to each analysis. Similarly, the reservoir and lines supplying deposition buffer to the valve and nozzle were flushed with 70% ethanol (EtOH), ddH2O, and 10 mM Tris-HCl at the beginning of each experiment. NaOH, ddH2O, and Tris-HCl were filter-sterilized in preparation. The electrophoretic background electrolyte and deposition buffer were matched (10 mM Tris-HCl, pH 7.5). Samples were injected for 0.5 s at 3.5 psi. Electrophoresis and fractionation began simultaneously. Page - 5


Page 6 of 26

Page 7 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Electrophoresis was performed at 233 V/cm, the deposition buffer was held under nitrogen pressure at 3.5 psi, and the nozzle was held at ground potential. The collection plate was secured to the motorized stage, which was programmed to move in a serpentine pattern: the plate moved to deposit a set of n fractions separated by a distance d in the X, then a distance d in the Y direction, followed by a row of n fractions deposited with distance d spacing in the -X direction, Figure 1.

Figure 1. Schematic of the capillary zone electrophoresis instrument coupled to a fraction collector. The distal end of the separation capillary is threaded through a Tee fitting and terminates at the exit of a precision dispensing nozzle. Sheath buffer is pressurized with nitrogen gas. Buffer flow is controlled with the dispensing valve to generate a drop that ensheaths the material exiting the capillary, depositing the drop onto a Petri dish. The Petri dish is mounted on a motorized microscope stage, which is programmed using Labview to move in a serpentine path; see supporting information for the Labview code that controls the instrument.

Page - 6



Instrument Sterilization: The system is contained within a custom enclosure, which reduces airflow and the opportunity to introduce environmental contamination. Methanol (MeOH, ≥ 99.9%, VWR) and 70% ethanol (EtOH) were routinely used on all surfaces. Capillary conditioning was modified to include routine flushing of MeOH followed by conditioning procedures onto a blank dish to ensure no viable microbial cells remained attached either inside or outside of the capillary and inlet. To ensure no viable microorganisms remained within the fractionation system, EtOH was added as a conditioning solvent in the deposition buffer reservoir; which sterilizes the internal nozzle surfaces. Sterility procedures were tested daily by electrophoretic fractionation of a blank injection of 10 mM Tris-HCl collected onto LB agar. The blank plates were incubated as described for experimental runs and absence of microbial growth was used to confirm sterilization. Culture-Dependent Analysis Fractions were deposited onto agar plates for analysis of culturable microbes. The motion of the stage was programmed in a 12x12 grid to match the dimensions of a standard Petri dish with 5 mm spot spacing (d = 5) in the X and Y dimensions. Fraction width, which controls time between depositions, was set to 8 s. Total fraction width, 9.3 s, includes stage travel time. Valve width, which controls the droplet volume, was programmed to dispense sub-microliter volumes of deposition buffer with each fraction. Fractionated E. coli bacteria were deposited onto LB agar supplemented with ampicillin and L-arabinose. Immediately after fractionation, plates were incubated at 37° C for 15 h. Plates were photographed under a UV lamp after incubation. Fractionated wastewater samples were

Page - 7


Page 8 of 26

Page 9 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


deposited onto LB agar or MacConkey agar (EMD, prepared according to manufacturer’s protocol), incubated at 32° C for 15 h, and photographed after incubation. Isolation, DNA Extraction, and Sequencing of Culturable Populations DNA was isolated for amplification of 16S rRNA DNA using colony boilates from streak purified colonies as described below. Boilates were prepared from the colonies formed in the culture-dependent analysis of wastewater microbiota: colonies from each fraction were picked and transferred to new LB-agar plates, re-growth was sampled and transferred to 100 μL sterile 1 x PBS, and heated to 95° C for 15 minutes to extract genomic DNA. Genomic DNA was clarified by centrifugation at 12,000 x g for 5 min. Samples were amplified using a CFX96 Touch Real-Time PCR Detection System (Bio-Rad) and universal 16S rRNA primers (ReadyMade Primers, IDT): forward primer 5’- AGA GTT TGA TCC TGG CTC AG, reverse primer 5’- ACG GCT ACC TTG TTA CGA CTT (29). The plates were sealed and centrifuged prior to real-time PCR: 95° C for 3 min followed by 40 cycles of 95° C for 10 s (denature), 55.8° C for 20 s (anneal), and 72° C for 20 s (extend). PCR products were purified (QIAquick PCR Purification Kit, Qiagen) and submitted for Sanger sequencing on the Applied Biosystems 96capillary model 3730xl DNA Analyzer (Genomics & Bioinformatics Core Facility at the University of Notre Dame). Sequences were analyzed using QIIME2 (version 2018.8, https://qiime2.org/) (30). Forward reads were imported for taxonomic assignment (see supporting document for sequences: Huge_16S_R1.fasta). Classification of bacterial 16S rRNA gene sequences was made using Silva (132 release) (31-32). The reference sequence database was pre-clustered at 99% ID, with amplicons for the domain of interest extracted using primer sequences with the featureclassifier’s extract_reads method (33-34).

Page - 8



RESULTS AND DISCUSSION Electrophoretic Fractionation of E. coli cells Capillary electrophoresis typically employs optical, mass spectrometric, or electrochemical detectors that are placed in-line or at the exit of the separation capillary. In this paper, we report the use of a Petri dish as a detector for culturable bacteria. This system is reminiscent of the radionucleotide detection in capillary electrophoresis reported by Tracht and colleagues; that system rastered the capillary tip across a membrane treated with a solid scintillator and integrated the scintillation with a CCD camera (35). In both Tracht’s and our cases, the electropherogram is revealed as a pattern along the serpentine path. In Tracht’s case, the signal is light emitted by the scintillator in response to radioactive decay. In our case, the signal is a set of colonies grown on the Petri dish. Unlike the scintillation-system, we employ a precision nozzle to deposit drops from the capillary, which eliminates contact with the surface and minimizes contamination. To visualize the performance of our system for a pure sample, plugs containing ~600 and 5,000 E. coli cells were injected into the separation capillary and subjected to electrophoresis. This E. coli strain expresses an arabinose-inducible GFP on a plasmid. Fractions were deposited in a serpentine pattern onto a Petri dish, incubated, and imaged under a UV lamp (Figure 2). Deposition began with the application of the electric field to the capillary.

Page - 9


Page 10 of 26

Page 11 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 2. Images of E. coli colonies after electrophoretic fractionation onto LB-agar prepared with ampicillin and L-arabinose. Left column: 5,000 cells were loaded by hydrodynamic injection. Right column: 600 cells were loaded by hydrodynamic injection. Reference grids are provided immediately to the right of each image to highlight the location of colony forming units (CFUs). Fraction width: 9.3 s. Fraction volume: 0.7 μL. Plates were incubated at 37° C for 15 hours and imaged under a UV lamp. In this system, the electropherogram is visualized as fluorescent colonies and demonstrates separation of intact organisms. Single colony counting is analogous to single molecule counting in molecular shot-noise limited experiments with fluorescence detection (36). Like single molecule detection, single bacteria detection is paralizable; however, colonies produced by co-deposition of more than one bacterium will merge into a single colony, which limits the dynamic range of the system. No colonies were observed until 7.3 minutes into the electrophoresis runs of Figure 2. During migration of this void volume, only sterile solution is deposited. At ~7.3 minutes, a set of Page - 10



fluorescent colonies was formed within the area defined by the deposited drop (E. coli migration time 7.4 ± 0.1 minutes, combined data). The first spot in each experiment is much more intense than subsequent spots, suggesting excellent electrophoretic separation efficiency. However, the very high amplification afforded by microbial incubation allows detection of individual bacteria that form a tailing peak. The two 600-cell injections (figure 2-right images) show no colonies after the initial spot. The first 5,000 cell injection (figure 2-top left image) shows approximately 12 colonies formed after the main spot. The second of the 5,000-cell injections (figure 2-bottom left image) shows evidence for more tailing. The use of a precision nozzle-based deposition system eliminates contact of the separation capillary with the surface of the Petri dish, which eliminates in situ contamination of the plate and nozzle. Preparative Electrophoretic Fractionation of Wastewater Microbiota After confirmation of cell viability and resolution following electrophoretic separation and deposition onto a Petri dish, we applied this technology to a complex environmental sample: microbiota isolated from the primary effluent of a regional wastewater treatment facility. After concentration and re-suspension in glycerol, serial dilutions were performed to inject 1,000 microbial cells into the capillary (as estimated by OD600). Fractions were deposited on LB plates in triplicate and MacConkey plates in duplicate. Figure 3 shows the colonies formed on LB agar after separation, collection, and incubation. Like analysis of E. coli, there is a void volume at the start of the run. An average of 15 colonies formed, which is consistent with the low culturable rate of environmental microbiota, the choice of media, and the growth conditions employed.

Page - 11


Page 12 of 26

Page 13 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Figure 3. Electrophoretic fractionation of a sewage microbiota obtained from primary effluent collected from the Mishawaka Indiana (USA) Wastewater Department; ~ 1,000 cells loaded by hydrodynamic injection, performed in triplicate. Fraction width: 9.3 s. Fraction volume: 0.35 μL. Plates were incubated at 32° C for 15 hours and photographed. Genomic DNA from each colony was extracted, the 16S rRNA gene was amplified, and the product was submitted for Sanger sequencing. The sequence data were used to identify each colony by matching forward reads to the Silva database. The colony marked Ø (A) was not identified. The gDNA extracted from this region did not amplify and was not sequenced. A deposition map below the images provides location information and corresponding family level taxa. Colonies were sampled for gDNA extraction. The 16S rRNA gene was amplified and sequenced by Sanger sequencing. Sequencing the 16S rRNA gene enables taxonomic assignment to each colony-producing fraction. The results of 16S sequencing are summarized in Table 1 (LB) and Table S1 (MacConkey). The microbial community is predominately Proteobacteria, which is common for wastewater effluent (37-39), with high abundance in Aeromonadaceae (37% Aeromonas sp.) and Moraxellaceae (24% Acinetobacter sp.) families.

Page - 12



The most common taxa observed are Aeromonas sp. The majority of the colonies identified as Aeromonas sp. (68%) consistently migrate at either 8.2 ± 0.08 minutes (position 5E) or 8.5 ± 0.08 minutes (position 5-G) on all LB plates. The remaining Aeromonas migrate independent of each other and span a separation window of nearly 12 minutes. A similar trend is observed for Acinetobacter sp., where there are reproducible migration patterns at positions corresponding to 8.7 ± 0.22 minutes and 10.9 ± 0.22 minutes. The remaining Acinetobacter span a separation window of 8.5 minutes. Additional instances of genus level reproducibility are found when these data are combined with the data presented in Figure S1 (taxon listed in Table S1), where electrophoretic fractionation was repeated onto selective media, MacConkey agar. An Acinetobacter sp. with migration time 8.2 ± 0.11 minutes is culturable on both medias (position 5-D on LB plate 2 and 5-E on MacConkey plate 1). Only two colonies were identified as Klebsiella sp., LB plate 3 at position 8-K and MacConkey plate 2 at position 8-L. These Klebsiella colonies have an average migration time of 13.3 ± 0.11 minutes. High dynamic range is common among environmental microbiomes. An important aspect of this work is the ability to fractionate the community so that abundant organisms (Aeromonas sp.) are segregated from the rare organisms to obtain greater depth and representation in analysis. In addition to the Klebsiella sp., there are two bacteria that are represented as isolated colonies: Micrococcus sp. (LB plate 1) and Uruburuella sp. (LB plate 3). Finally, a group of colonies on LB plate 1 at position 7-E were identified as Citrobacter sp. with a migration time of 12.0 minutes.

Page - 13


Page 14 of 26

Page 15 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


CONCLUSION We demonstrate the use of capillary zone electrophoresis separation, followed by deposition of fractions onto a Petri dish coupled with Sanger sequencing of a phylogenetic marker, for identification of culturable microbes within complex microbiomes. Since a single bacterium can form a colony on the dish, this system has unrivaled sensitivity. The 16S rRNA sequence has been tabulated for a large number of species, and provides the starting point for characterization of the microbes in a microbiome. However, based on the 16S rRNA sequence, we are not able to account for species and strain level differences, the presence of particular genes, or other forms of phenotypic heterogeneity that may perturb the charge or size of the bacteria (40). Our technology has one obvious limitation: it can only study culturable bacteria. Only a few percent of microbes in an environmental microbiome is culturable, so that a large population of microbes is invisible to analysis. As a less obvious limitation, sampling inevitably limits the precision and accuracy of our characterization of this very complex system (41). There is clearly a trade-off between our ability to resolve colonies and the depth to which we can probe the microbiome. Finally, it has not escaped our attention that this technology will be particularly useful for metagenomic analysis of the microbiome. In metagenomics, DNA from the microbiome is sheared randomly, and the resulting fragments are sequenced using next-generation sequencing technology. Conventional metagenomic studies are plagued by tremendous redundancy, where highly abundant species generate a very large fraction of the sequencing reads and where assembly of the sequences into a finished genome is plagued by interferences from the sequences

Page - 14



of closely related species. Separation of the microbiome into simpler fractions should significantly reduce the challenges associated with redundant sequences.

Page - 15


Page 16 of 26

Page 17 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


ASSOCIATED CONTENT Supporting Information Labview block diagrams with file descriptions Sanger sequence forward reads: Huge_16S_R1.fasta Electrophoretic fractionation of wastewater microbiota collected onto MacConkey agar Taxonomic assignments for colonies formed on MacConkey agar

ACKNOWLEDGMENTS We acknowledge funding from the Advanced Diagnostics and Therapeutics program at the University of Notre Dame (NJD & MMC), a NSF graduate fellowship (BJH), and the Walton Family Foundation 2017-1936 (MMC & NJD). The Labview program controlling fraction collection has evolved over a decade’s period by members of the Dovichi Laboratory. We particularly thank Michael Vannatta, Colin Whitmore, Oluwatosin Dada, and Scott Sarver for their contributions to the development of this software. We also thank Andrew Schmudlach for his contributions to buffer development. We thank the John R. Kirby Lab of the Medical College of Wisconsin for helpful discussions. We particularly thank Orlando deLeon for assistance with data processing. We also acknowledge the assistance of the Genomics facility at Notre Dame for 16S sequencing. We also acknowledge the assistance of the staff at the Mishawaka, IN (USA) Wastewater Department for sample collection. This material is based upon work supported by the National Science Foundation Graduate Research Fellowship under Grant no. DGE-1313583. Any opinion, findings, and conclusions or

Page - 16



recommendations expressed in this material are those of the authors(s) and do not necessarily reflect the views of the National Science Foundation.

Conflict of interest statement. A patent application has been filed by the University of Notre Dame for the technology described in this manuscript; that application lists this paper’s authors as inventors.

Page - 17


Page 18 of 26

Page 19 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


REFERENCES 1.

Staley, J. T.; Konopka, A. Measurement of in situ Activities of Nonphotosynthetic Microorganisms in Aquatic and Terrestrial Habitats. Annu. Rev. Microbiol. 1985, 39, 321–346.

2.

Stewart, E. J. Growing Unculturable Bacteria. J. Bacteriol. 2012, 194, 4151–4160.

3.

Bechhold, H. Coagulation of Suspensions or Colloids and Bacterial Agglutination. Phys. Chem. 1904, 48, 385–423.

4.

Brown, H. C.; Broom, J. C. Further Observations on Bacterial Cataphoresis. Br. J. Exp. Pathol. 1932, 13, 334–337.

5.

Moyer, L. S. Changes in the Electrokinetic Potential of Bacteria at Various Phases of the Culture Cycle. J. Bacteriol. 1936, 32, 433–464.

6.

Dyar, M. T.; Ordal, E. J. Electrokinetic Studies on Bacterial Surfaces: I. Effects of Surface-Active Agents on Electrophoretic Mobilities of Bacteria. J. Bacteriol. 1946, 51, 149–167.

7.

Hartman, R. S.; Eigelsbach, H. T.; Bateman, J. B.; Braun, W.; Herring, R.; Rodgers, R. D. An Electrophoretic Method for the Assay of Bacterial Variants. Appl. Microbiol. 1953, 1, 178–183.

8.

Bayne, D. R.; Lawrence, J. M. Separating Constituents of Natural Phytoplankton Populations by Continuous Particle Electrophoresis. Limnol. Ocean. 1972, 17, 481–489.

9.

Schott, H.; Young, C. Y. Electrokinetic Studies of Bacteria I: Effect of Nature, Ionic Strength, and PH of Buffer Solutions on Electrophoretic Mobility of Streptococcus faecalis and Escherichia coli. J. Pharm. Sci. 1972, 61, 182–187.

10.

Schott, H. The Effect of Buffers and of Organic and Inorganic Cations on the

Page - 18



Electrokinetic Properties of Bacteria. Bioelectrochemistry Bioenerg. 1977, 4, 117–136. 11.

Lytle, D. A.; Johnson, C. H.; Rice, E. W. A Systematic Comparison of the Electrokinetic Properties of Environmentally Important Microorganisms in Water. Colloids Surfaces B Biointerfaces 2002, 24, 91–101.

12.

Sloyer, J. L.; Bayer, M. E. Automated Electrokinetic Analysis; Description and Application in Virology and Cell Biology. J. Virol. Methods 1990, 28, 207–216.

13.

Ebersole, R. C.; McCormick, R. M. Separation and Isolation of Viable Bacteria by Capillary Zone Electrophoresis. Biotechnology. (N. Y). 1993, 11, 1278–1282.

14.

Armstrong, D. W.; Schulte, G.; Schneiderheinze, J. M.; Westenberg, D. J. Separating Microbes in the Manner of Molecules. 1. Capillary Electrokinetic Approaches. Anal. Chem. 1999, 71, 5465–5469.

15.

Dai, D.; Chen, Y.; Qi, L.; Yu, X. Critical Factors for Reproducible Capillary Electrophoresis of Escherichia coli Cells. Electrophoresis 2003, 24, 3219–3223.

16.

Yu, L. J.; Li, S. F. Y. Electrophoretic Behavior Study of Bacteria of Pseudomonas Aeruginosa, Edwardsiella Tarda and Enteropathogenic Escherichia Coli by Capillary Electrophoresis with UV and Fluorescence Detection. Chromatographia 2005, 62, 401– 407.

17.

Bao, Y.; Lantz, A. W.; Crank, J. A.; Huang, J.; Armstrong, D. W. The Use of Cationic Surfactants and Ionic Liquids in the Detection of Microbial Contamination by Capillary Electrophoresis. Electrophoresis 2008, 29, 2587–2592.

18.

Lantz, A. W.; Brehm-Stecher, B. F.; Armstrong, D. W. Combined Capillary Electrophoresis and DNA‐fluorescence in Situ Hybridization for Rapid Molecular Identification of Salmonella Typhimurium in Mixed Culture. Electrophoresis 2008, 29,

Page - 19


Page 20 of 26

Page 21 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


2477–2484. 19.

Haugg, M.; Kaiser, V.; Schmidtkunz, C.; Welsch, T. The Effect of Aggregation on the Separation Performance of Bacteria in Capillary Electrophoresis. Electrophoresis 2009, 30, 396–402.

20.

Soykut, E. A.; Boyaci, I. H. Analyzing and Monitoring of Phage-Bacteria Interaction Using CE. Electrophoresis 2009, 30, 3548–3554.

21.

Tong, M.-Y.; Jiang, C.; Armstrong, D. W. Fast Detection of Candida Albicans and/or Bacteria in Blood Plasma by “Sample-Self-Focusing” Using Capillary ElectrophoresisLaser-Induced Fluorescence. J. Pharm. Biomed. Anal. 2010, 53, 75–80.

22.

Horká, M.; Růžička, F.; Kubesová, A.; Němcová, E.; Šlais, K. Separation of Phenotypically Indistinguishable Candida Species, C. Orthopsilosis, C. Metapsilosis and C. Parapsilosis, by Capillary Electromigration Techniques. J. Chromatogr. A 2011, 1218, 3900–3907.

23.

Szeliga, J.; Jackowski, M.; Kłodzińska, E.; Buszewski, B.; Kupczyk, W. Clinical Application of a Rapid Microbiological Test Based on Capillary Zone Electrophoresis to Assess Local Skin Infection. BMC Res. Notes 2011, 4, 467.

24.

Szumski, M.; Kłodzińska, E.; Dziubakiewicz, E.; Hrynkiewicz, K.; Buszewski, B. Effect of Applied Voltage on Viability of Bacteria during Separation under Electrophoretic Conditions. J. Liq. Chromatogr. Relat. Technol. 2011, 34, 2689–2698.

25.

Dziubakiewicz, E.; Buszewski, B. Capillary Electrophoresis of Microbial Aggregates. Electrophoresis 2014, 35, 1160–1164.

26.

Krylov, S. N.; Starke, D. A.; Arriaga, E. A.; Zhang, Z.; Chan, N. W.; Palcic, M. M.; Dovichi, N. J. Instrumentation for Chemical Cytometry. Anal. Chem. 2000, 72, 872–877.

Page - 20



27.

Vannatta, M. W.; Whitmore, C. D.; Dovichi, N. J. CE-MALDI Interface Based on Inkjet Technology. Electrophoresis 2009, 30, 4071–4074.

28.

Huge, B. J.; Flaherty, R. J.; Dada, O. O.; Dovichi, N. J. Capillary Electrophoresis Coupled with Automated Fraction Collection. Talanta 2014, 130, 288–293.

29.

Weisburg, W. G.; Barns, S. M.; Pelletier, D. A.; Lane, D. J. 16S ribosomal DNA amplification for phylogenetic study. Journal of Bacteriology 1991, 173, 697-703.

30.

Caporaso, J. G.; Kuczynski, J.; Stombaugh, J.; Bittinger, K.; Bushman, F. D.; Costello, E. K.; Fierer, N.; Peña, A. G.; Goodrich, J. K.; Gordon, J. I.; et al. QIIME Allows Analysis of High-Throughput Community Sequencing Data. Nat. Methods 2010, 7, 335–336.

31.

Quast, C.; Pruesse, E.; Yilmaz, P.; Gerken, J.; Schweer, T.; Yarza, P.; Peplies, J.; Glöckner, F. O. The SILVA Ribosomal RNA Gene Database Project: Improved Data Processing and Web-Based Tools. Nucleic Acids Res. 2012, 41, D590–D596.

32.

Yilmaz, P.; Parfrey, L. W.; Yarza, P.; Gerken, J.; Pruesse, E.; Quast, C.; Schweer, T.; Peplies, J.; Ludwig, W.; Glöckner, F. O. The SILVA and “All-Species Living Tree Project (LTP)” Taxonomic Frameworks. Nucleic Acids Res. 2014, 42, D643–D648.

33.

Werner, J. J.; Koren, O.; Hugenholtz, P.; DeSantis, T. Z.; Walters, W. A.; Caporaso, J. G.; Angenent, L. T.; Knight, R.; Ley, R. E. Impact of Training Sets on Classification of High-Throughput Bacterial 16S rRNA Gene Surveys. ISME J. 2012, 6, 94–103.

34.

Bokulich, N. A.; Kaehler, B. D.; Rideout, J. R.; Dillon, M.; Bolyen, E.; Knight, R.; Huttley, G. A.; Gregory Caporaso, J. Optimizing Taxonomic Classification of MarkerGene Amplicon Sequences with QIIME 2’s Q2-Feature-Classifier Plugin. Microbiome 2018, 6, 90.

35.

Tracht, S.; Toma, V.; Sweedler, J. V. Postcolumn Radionuclide Detection of Low-Energy

Page - 21


Page 22 of 26

Page 23 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


Beta Emitters in Capillary Electrophoresis. Anal. Chem. 1994, 66, 2382–2389. 36.

Chen, D.; Dovichi, N. J. Single-Molecule Detection in Capillary Electrophoresis: Molecular Shot Noise as a Fundamental Limit to Chemical Analysis. Anal. Chem. 1996, 68, 690–696.

37.

McLellan, S. L.; Huse, S. M.; Mueller-Spitz, S. R.; Andreishcheva, E. N.; Sogin, M. L. Diversity and Population Structure of Sewage-Derived Microorganisms in Wastewater Treatment Plant Influent. Environ. Microbiol. 2010, 12, 378–392.

38.

Ye, L.; Zhang, T. Bacterial Communities in Different Sections of a Municipal Wastewater Treatment Plant Revealed by 16S RDNA 454 Pyrosequencing. Appl. Microbiol. Biotechnol. 2013, 97, 2681–2690.

39.

Cydzik-Kwiatkowska, A.; Zielin, M. Bacterial Communities in Full-Scale Wastewater Treatment Systems. World J Microbiol Biotechnol 2016, 32, 1–8.

40.

Ackermann, M. A Functional Perspective on Phenotypic Heterogeneity in Microorganisms. Nat. Rev. Microbiol. 2015, 13, 497–508.

41.

Kratochvil, B.; Taylor, J.K. Sampling for chemical analysis. Anal. Chem. 1981, 53, A924-A938.

Page - 22



Page 24 of 26

Table 1. Taxonomic Assignments for Culturable Wastewater Microbiota using the SILVA Database CFU

Grid

1 2

5-E 5-E

3

5-F

4

5-G

5

5-G

6

5-G

7

5-L

8

6-I

9

6-B

10 11

7-C 7-E

12

7-E

13

7-K

14

8-J

15 16

8-H 8-G

17 18

8-E 8-D

19

9-E

1

5-D

2

5-E

3

5-G

4

5-G

5

5-I

6

6-D

7

7-F

Taxon Plate 1 Bacteria Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas hydrophila subsp. hydrophila Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae; Citrobacter Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae; Citrobacter Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae; Enterobacter Bacteria Bacteria; Actinobacteria; Actinobacteria; Micrococcales; Micrococcaceae; Micrococcus Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Plate 2 Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter sp. WCHA60 Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter

Page - 23


Confidence 1.00 0.94 1.00 1.00 0.74 1.00 1.00 1.00 1.00 1.00 0.75 0.84 1.00 0.98 0.99 0.96 1.00 1.00 1.00 1.00 1.00 1.00 1.00 0.95 1.00 1.00

Page 25 of 26 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60


CFU 8

Grid 8-C

9

9-B

10

11-D

1

5-C

2

5-C

3

5-C

4

5-D

5

5-D

6

5-E

7

5-E

8

5-F

9

5-F

10

5-G

11

5-J

12

7-H

13

8-K

14

10-L

15

12-L

16

12-I

Taxon Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas hydrophila subsp. hydrophila Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Plate 3 Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas Bacteria; Proteobacteria; Gammaproteobacteria; Enterobacteriales; Enterobacteriaceae; Klebsiella Bacteria; Proteobacteria; Gammaproteobacteria; Betaproteobacteriales; Neisseriaceae; Uruburuella metagenome Bacteria; Proteobacteria; Gammaproteobacteria; Pseudomonadales; Moraxellaceae; Acinetobacter Bacteria; Proteobacteria; Gammaproteobacteria; Aeromonadales; Aeromonadaceae; Aeromonas

Page - 24


Confidence 0.72 1.00 1.00 1.00 0.99 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 1.00 0.96 1.00 1.00 1.00


Table of content graphic

Page - 25


Page 26 of 26

Capillary zone electrophoresis with fraction collection for separation

Recommend Documents