16.2: Micro-report 1- Selective plating experiment - Biology

16.2: Micro-report 1- Selective plating experiment - Biology

We are searching data for your request:

Forums and discussions:
Manuals and reference books:
Data from registers:
Wait the end of the search in all databases.
Upon completion, a link will appear to access the found materials.

The figure is at the heart of every micro-report. This multi-panel figure of a spot plating experiment was prepared by students in an advanced lab class.

Note the following features of the figure:

  • Plates are oriented in the same direc- tion so they can be easily compared
  • Strains and media are labeled. Strain names can be used in the figure itself or indicated by a code that is defined in the legend.
  • Strain names are used when the genotypes of strains are uncertain (your experiment). This figure presents results from an experiment where genotypes were known.
  • The reader will need to refer to the M&M section for additional details about the media and experiment.

Specific guidelines for micro-report 1 follow.

Purpose: You have three yeast strains derived from strain, BY4742. In one sentence, what are you trying to do in this experiment?

Materials and Methods: In preparing the M&M, ask yourself “What information will an investi- gator need to reproduce our experiments?” Provide information on the strains and media that you used, as well as the procedures that you used for spot plating.

Strains: Include the names of your strains as well as the genotype of the BY4742 parent strain.

Media: Identify the culture media you used in the experiments. Decide on a naming conven- tion - the same nomenclature should be used in both the figure and M&M. Reference the manual for the composition of the media, rather than including all the components here.

Spot plating: Someone trying to reproduce your results will need to know how your starter cul- tures were generated (cultures were grown overnight in YPD) and how cultures were diluted for the spot plates (e.g. a series of 1:10 dilutions in sterile water). They do NOT need to know that you transferred 10 μL yeast culture to 90 μL water. Readers will need to know how many microliters were used for each spot and the conditions used to incubate (time, temperature) the plates.

Results and Discussion - Your figure with the scanned plates is the focal point of this section. The R&D section tells a story of how you used your plating data to identify the met deletions in the YMP strains. Provide a brief narrative that guides your reader through your results and your thinking. How would deletions in your MET genes affect the ability of your YMP strains to use different sulfur sources? Do the data allow you to confidently identify the strains?

A single summary data table documenting the growth of YMP and BY4742 strains on various culture media is a good way to bring together the experimental data and your conclusions. Describe the growth of the met deletion strains on the various media and include your preliminary strain identifications from the experimental data.

Metagenomic engineering of the mammalian gut microbiome in situ

Engineering of microbial communities in open environments remains challenging. Here we describe a platform used to identify and modify genetically tractable mammalian microbiota by engineering community-wide horizontal gene transfer events in situ. With this approach, we demonstrate that diverse taxa in the mouse gut microbiome can be modified directly with a desired genetic payload. In situ microbiome engineering in living animals allows novel capabilities to be introduced into established communities in their native milieu.


The abundance of genetic abnormalities and phenotypic heterogeneities in AML pose significant challenges to developing improved treatments. Here we demonstrated that a key GAS6/AXL axis is highly activated in AML patient cells, particularly in stem/progenitor cells. We developed a potent, selective AXL inhibitor that has favorable pharmaceutical properties and efficacy against preclinical patient-derived xenotransplantation (PDX) models of AML. Importantly, inhibition of AXL sensitized AML stem/progenitor cells to venetoclax treatment, with strong synergistic effects in vitro and in PDX models. Mechanistically, single-cell RNA-sequencing and functional validation studies uncovered that AXL inhibition or in combination with venetoclax potentially targets intrinsic metabolic vulnerabilities of AML stem/progenitor cells, which shows a distinct transcriptomic profile and inhibits mitochondrial oxidative phosphorylation. Inhibition of AXL or BCL-2 also differentially targets key signaling proteins to synergize in leukemic cell killing. These findings have direct translational impact on the treatment of AML and other cancers with high AXL activity.


Transgenic plants: survival and EcoRI inheritance and expression

Initially, nine putative independent transgenic tobacco (Nicotiana tabacum cv. Xanthi) events were generated that harboured the EcoRI construct (Figure 1a) lines were downselected as described below. Southern blot analysis revealed that transgenic lines E1, E2, E3 and E4 were positive for the hygromycin gene, and lines E7, E8 and E9 had faint bands of high molecular weight (Figure S1). No bands were observed in the blot for the nontransgenic control tobacco. Line E6 did not survive when grown in the glasshouse, and line E5 apparently did not express the transgene these two lines were excluded in subsequent analyses. All remaining T0 EcoRI events produced were self-pollinated to produce T1- and subsequently T2-progeny seeds. Each T0 event was expected to be hemizygous at each insertion locus, and the maintenance of hemizygotes and homozygous nulls would be expected for each generation if the transgene product ablated pollen. If EcoRI effectively ablated pollen, the transmission of the hpt gene (hygromycin phosphotransferase, conferring resistance to hygromycin) would be prevented, and the expected segregation ratio for a single-copy selfed line would be 1 : 1 transgenic : nontransgenic). However, transgenic progeny percentage could be lower if the female gametophyte or seed development is affected by EcoRI expression or higher if the transgene copy number was greater than one.

To test for the predicted segregation ratio, progeny analysis on selective media showed that T2 generation progeny significantly deviated from the expected 1 : 1 ratio of hygromycin-resistant (hmR) to hygromycin-susceptible (hmS) individuals with the exception of line E3 (Table S1). The transmission of hmR was reduced through the female for lines E1, E3 and E4 as determined by significantly fewer hmR individuals (≤1 : 1) than predicted. Further, lines E7, E8 and E9 demonstrated increased hmR in the populations as the observed ratios were ≥1 : 1 for hmR to hmS. All nontransgenic seedlings died when grown on selective media (Table S1), and all seedlings survived when grown on nonselective media (data not shown).

In pollen, the EcoRI transcript was present in all transgenic lines with none detected in nontransgenic control plants (Figures 2a and S2). Line E1 had the highest EcoRI transcript abundance than other transgenic lines (Figure 2a), and line E9 had the lowest transcript abundance (Figure S2). Lines E7, E8 and E9 were eliminated from further tissue-specific EcoRI transcript analysis owing to progeny analysis results from crossing experiments. For all other tissues analysed, there was negligible EcoRI transcript detected (Figure 2b). The EcoRI transcript abundance was <0.5% of the lowest signal observed in pollen. However, transcript abundance in stems from line E3 was significantly higher than all others (Figure 2b). In the case of line E3, there was large variation in measurements among biological replicates (replicate 1 = 0.01, replicate 2 = 0.14, and replicate 3 = 0.00), but no morphological variation was observed.

Plant morphology

Transgenic plants were morphologically indiscernible from nontransgenic control plants (Figure 3), with the exception of less biomass produced by transgenic lines E1, E4 and E7 (P ≤ 0.01) (Table 1). The biomass production and stem circumference of lines E1 and E4 were significantly lower than all others (Table 1). Transgenic plant height (P = 0.66), internode length (P = 0.10) and number of flowers per inflorescence (P = 0.50) were not significantly different among transgenic lines and the control (Table 1).

Line N Dry total biomass (g) Plant height (cm) Stem circumference 5 cm above soil level (cm) Internode length (cm) Flower number per inflorescence
Nontransgenic 5 541.2 ± 27.8 a 146.5 ± 0.8 5.3 ± 0.1 a 6.2 ± 0.4 28 ± 3
E1 5 394.4 ± 21.5 c 140.3 ± 1.0 4.6 ± 0.1 c 6.4 ± 0.5 28 ± 3
E2 5 529.3 ± 60.0 ab 146.0 ± 1.0 5.0 ± 0.04 ab 7.3 ± 0.6 31 ± 3
E3 5 528.0 ± 17.0 ab 145.7 ± 2.5 5.1 ± 0.1 a 7.7 ± 0.3 27 ± 2
E4 5 382.2 ± 31.0 c 148.4 ± 0.7 4.7 ± 0.0 bc 7.2 ± 0.8 27 ± 3
E7 5 434.6 ± 30.3 bc 145.9 ± 1.8 5.0 ± 0.1 abc 6.6 ± 0.5 26 ± 3
E8 5 449.6 ± 25.6 abc 143.5 ± 1.9 5.0 ± 0.0 ab 7.7 ± 0.5 32 ± 4
E9 5 516.4 ± 44.6 ab 137.4 ± 0.5 5.0 ± 0.0 abc 5.7 ± 0.5 31 ± 3
  • Superscript letters within a column represent significant differences (Fisher's LSD, P ≤ 0.05).

Pollen counts were similar among transgenic lines and the nontransgenic control (Figure S3). Several glasshouse-grown transgenic lines produced significantly fewer seeds per pod compared with the control (Figure 4a, P = 0.03), with no differences evident among lines E4, E7 and the control. When grown under field conditions, the mean seed production was apparently lower for lines E1 and E4, but no significant differences were detected among lines and the control (Figure 4b).


When hand-crossed with male-sterile stamenless tobacco (cv. ‘MS TN90’) in the glasshouse, all transgenic lines we assayed produced F1 seed, and germination was apparently normal in the glasshouse and field crosses (Table 2). The crosses using lines E1, E2 and E3 did not produce any transgenic progeny in the glasshouse, resulting in 100% efficient bioconfinement, whereas other lines produced at least one transgenic seed (Table 2). Line E4 produced the most transgenic progeny followed by lines E8 and E7 however, all progeny produced from transgenic lines were >99% transgene-free. All transgenic progeny that survived hygromycin selection (Figure S4) appeared to be morphologically normal. Additionally, all transgenic lines had decreased pollen germination compared with the nontransgenic control (P ≤ 0.01 Table 2). Line E4 had the highest rate of pollen germination among all the transgenic lines.

Line Bioconfinement efficiency (%) Transgenic progeny Seeds germinated Seeds screened Pollen germination (%)
Glasshouse MS TN90 cross -progeny screen
Nontransgenic N/A 0 36 860 38 800 42.33 ± 0.08 a
E1 100.00 0 38 000 40 000 18.33 ± 0.09 c
E2 100.00 0 28 310 29 800 21.25 ± 0.09 bc
E3 100.00 0 38 000 40 000 19.75 ± 0.04 bc
E4 99.12 248 38 000 40 000 30.58 ± 0.04 b
E7 99.99 1 38 000 40 000 17.58 ± 0.04 c
E8 99.77 36 22 420 23 600 17.33 ± 0.05 c
E9 99.85 17 15 200 16 000 12.50 ± 0.05 c
Field MS TN90 cross -progeny screen
Nontransgenic N/A 0 8124 9249 38.51 ± 0.05
E1 100.00 0 1690 1764 28.34 ± 0.02
E2 99.95 1 2037 2167 27.94 ± 0.03
E3 100.00 0 861 909 34.26 ± 0.04
E4 100.00 0 1021 1253 30.73 ± 0.02
  • Superscript letters within a column represent significant differences (Fisher's LSD, P ≤ 0.05).

All open-pollinated field experiments resulted in F1 seed collected from MS TN90. As hand-crossing was not performed in the field, the total seed number screened for transgenicity was noticeably lower than crosses performed in the glasshouse experiments (Table 2). Of the 13 733 F1 progeny seedlings screened on hygromycin media, a single surviving transgenic F1 progeny was detected from line E2. Seeds produced in plots from lines E1, E2 and E4 were 100% transgene-free (Table 2). When compared with controls, pollen germination means were slightly lower for field-grown transgenic lines, but no significant differences were detected (P = 0.15 Table 2).

Similar results were observed when transgenic lines were crossed to emasculated nontransgenic tobacco (cv. ‘Xanthi’) over multiple generations. Lines E1, E2 and E3 produced no transgenic F1 progeny, again, resulting in 100% bioconfinement (Table S2). Lines E4, E7, E8 and E9 produced transgenic progeny, but all F1 populations were 99% transgene-free (Table S2). Transgenic F1 individual plants from lines E4, E7, E8 and E9 were again backcrossed to emasculated ‘Xanthi’ to produce BC1F1 progeny. Bioconfinement efficiencies were recorded for each plant within a transgenic line and ranged from 99% to 100% in all BC1F1 progeny populations (Table S2). Backcrosses performed with F1 individuals from line E7 produced no BC1F1 seed.

Virtual Lab Experiments in Biotechnology: Bacterial Transformation

Cold Spring Harbor Laboratory’s DNA Learning Center presented this course as a service to help engage teachers and students in China during the coronavirus school closures.

The bacterial transformation experiment illustrates the direct link between an organism's genetic complement (genotype) and its observable characteristics (phenotype). A gene for antibiotic resistance is introduced into the bacterium E. coli. Following overnight incubation, transformed bacteria are compared to unexposed bacteria for their ability to grow in the presence of ampicillin.

Duration: 1 hour, 4 minutes, 4 seconds

e coli bacteria,dna transformation,dna molecule,herbert boyer,stanley cohen,recombinant dna,dna sequence,e coli,plasmid,expression

Related Content

15916. DNA transformation

Stanley Cohen and Herbert Boyer inserted the recombinant DNA molecule they created into E. coli bacteria by means of a plasmid, thereby inducing the uptake and expression of a foreign DNA sequence known as "transformation."

16705. Animation 34: Genes can be moved between species.

Stanley Cohen and Herbert Boyer transform bacteria with a recombinant plasmid, and Doug Hanahan studies induced transformation.

16721. Biography 34: Herb W. Boyer (1936 - )

Herb Boyer and Stan Cohen "invented" recombinant DNA technology.

15074. Taking apart plasmid DNA, Stanley Cohen

Stanley Cohen speaks about his and Herbert Boyer's experiment to make the first plasmid that had been engineered to contain foreign DNA.

15918. Transformation

DNA transformation is a naturally occuring but rare event in which DNA can be transferred into bacteria. In 1970, Morton Mandel and Akiko Higa discovered a way to make E. coli more "competent" for transforming foreign DNA. Their calcium chloride method is

15162. Preparing the DNA for sequencing, Shane Yeager

Shane Yeager, from the Whitehead Institute Center for Genome Research, explains the processes of storing and preparing DNA for sequencing.

15028. Interest in plasmids, Herbert Boyer

Herb Boyer talks about Stanley Cohen's and his interest in plasmids as vectors for DNA.

15915. The first recombinant DNA

Stanley Cohen and Herbert Boyer's historic experiment used techniques to cut and paste DNA to create the first custom-made organism containing recombined or "recombinant" DNA.

15020. Possible dangers of recombinant DNA, Paul Berg

Paul Berg talks about possible dangers of recombinant DNA.

15626. Paul Berg and Stanley Cohen

Herbert Boyer: Former varsity lineman turned biotech bigwig. Expert at cutting DNA before most people knew it could be done. Stanley Cohen: A born tinkerer figured out the trick of using loops of DNA called plasmids to transform bacterial DNA

Nematode and bacterial strains

C. elegans N2 (ancestral) strain used for all experiments unless otherwise indicated was kindly provided by J.J. Ewbank (Marseille, France). Strain TJ375 (gpIs1[hsp-16.2::GFP]) and mutant strains CF1038 (daf-16 (mu86) I), GR1307 (daf-16 (mgDf50) I), NU3 (dbl-1 (nk3) V), and KU25 (pmk-1(km25) IV) were obtained from the Caenorhabditis Genetics Center (Minneapolis, Minnesota, USA). N2 control was included in every experiment with nematode mutants.

E. coli strains used in this study were pathogenic strain 536 [23], and the uracil deficient strain OP50 [16]. Genotypic and phenotypic characterization of these strains was described previously [19]. Enterococcus faecalis strain OG1RF [63] (previously named Streptococcus faecalis OG1-RF1) was kindly provided by Dr. P. Courvalin (Paris, France).

Nematode maintenance and synchronization

Nematodes were maintained at 25°C on nematode growth medium (NGM) agar plates seeded with 0.1 mL LB grown stationary phase bacterial culture, and incubated 18 h at 37°C to densities of 7.3 × 10 9 ± 3 × 10 8 , 1.4 × 10 10 ± 5 × 10 9 , and 1.5 × 10 10 ± 2 × 10 9 CFU/plate for E. coli OP50, 536, and E. faecalis OG1RF, respectively. Age-synchronized populations of nematodes were initiated from eggs recovered following sodium hydroxide (0.5 M final) and sodium hypochlorite (0.96% final) treatment of gravid adults maintained at 25°C and fed E. coli OP50. All assays were carried out at 25°C with nematodes synchronized a second time at the end of development by selecting exclusively nematodes at the end of the 4 th larval (L4) stage based on vulva morphology.

Survival assays

Survival assays were carried out at least in triplicate. For survival assays, all nematode strains were developed and maintained at 25°C, transferred onto new plates every day during the first 5 days to eliminate progeny, and every 2–3 days thereafter. Dead nematodes were scored every 24 h. A nematode was considered dead when it failed to move spontaneously or respond to a gentle touch with a platinum wire. Nematodes buried in the agar or on the sides of the plates were censured from the analyses. Lifespan was measured as the time from the end of L4 larval stage (beginning of adulthood) until death.

Elimination of bacteria from nematodes intestine

When experiments involved nematodes developed on E. coli 536 or E. faecalis OG1RF and then transfer to a different bacteria strain, L4 stage nematodes were washed and treated with antibiotics to remove the initial strain from their intestinal tract. Nematodes were washed in M9 minimal salts buffer to remove excess bacteria from their surface and then incubated in NGM for 1 h to allow them to expurgate intestinal bacteria. Nematodes were then treated for 1 h with 20 μg/ml Polymyxin B antibiotic in M9 minimal salts buffer. Finally, nematodes were washed in M9 minimal salts buffer and transferred on plates containing the appropriate bacterial strain. The effect of antibiotic treatment on nematode survival was taken into account by applying the treatment to all nematode populations involved in the assays. For survival experiments, treated nematodes were regularly checked for the absence of the initial bacterial strain. A few nematodes were sampled from the survival assay, crushed in a Dounce homogenizer and the extract plated on LB agar media for colony observation (see below quantification of live bacterial cells in nematode intestine for detail on this procedure). Colony morphology and color allowed for direct visual discrimination between the different bacterial strains used in this study.

Measurements of nematode heat shock resistance

L4 nematodes developed on OP50 or 536 were washed in Polymyxin B antibiotic (see above elimination of bacteria from nematodes intestine). Then, nematodes were transferred onto OP50 lawn in order to measure heat shock resistance using nematodes that have the same bacterial strain in their intestines. After allowing nematodes to recover for 12 h at 25°C, they were transferred for 10 h at 35°C. Dead nematodes were scored at the end of the 10 h incubation at 35°C. This assay was performed in triplicate with internal triplicate controls.

Measurements of nematode heat shock protein (HSP) expression

HSP expression measurement assays were carried out with C. elegans strain TJ375 carrying a fluorescent reporter under the control of the hsp-16.2 promoter (gpIs1hsp-16-2::GFP]). Nematodes were collected with cold (4°C) sterile water, fixed immediately by addition of an equal volume of cold 2% formaldehyde in 2 × Phosphate Buffered Saline (PBS), and incubated 10 min on ice. Fixed nematodes were collected by gravity sedimentation and washed in cold 1× PBS before being loaded into a 96-well plate kept at 4°C in the dark until analyzed. Two-day old adults were separated from their progeny on the first and second day by gravity sedimentation in 15 ml falcon tubes for 2 min in M9 minimal salts buffer, the larvae being removed with the supernatant. The absence of eggs and larvae was verified under a dissecting microscope. Gene reporter levels were quantified with the COPAS Biosort (Union Biometrica) as described in [64]. Briefly, worms were analyzed for size (TOF) and green (GFP) fluorescence. Raw data were filtered on the TOF (200–1000) to exclude dust, bubbles, and aggregated worms. Fluorescence data was acquired from two independent experiments, including internal replicates.

Quantification of live bacterial cells in nematode intestine

Two-day old nematodes were handled at 4°C to stop defecation. Nematodes were transferred to a new plate without bacteria, recovered in 1.5 ml 10 -2 M MgSO4, and vortexed 30 seconds to remove excess bacteria from nematode cuticles. Then, individual nematodes were again transferred to a new plate without bacteria and rubbed across the plate to remove all bacteria from the cuticle. Nematodes were then individually crushed in a Dounce homogeneizer with the fitted mortar (B) in 10 -2 M MgSO4. The amount of live bacteria was determined by plating of appropriate dilutions on LB agar. After overnight incubation at 37°C, grown colonies were counted.

Statistical analyses and figures

Statistical analyses and graphic displays were made using Prism 5.0d from GraphPad Software, Inc. Measurement assays were analyzed by unpaired t-test for comparison of two groups or, when more than two groups were involved, using 1-way analysis of variance (ANOVA) followed by Tukey’s Multiple Comparison Test (TMCT) also known as Tukey-Kramer test, comparing all pairs of group and allowing for unequal sample sizes. Survival assays were analyzed using the Gehan-Breslow-Wilcoxon Test (GBWT) comparing conditions by pairs and allowing for unequal hazard ratios. Data presented are mean ± s.e.m., unless otherwise indicated. For survival assays, a typical, representative experiment is presented, the absence of significant difference between replicates was verified using GBWT .

Materials and Methods

Experimental model

HeLa cells and HeLa Kyoto BAC cell lines were grown at 37°C and 5% CO2 in DMEM high glucose (4.5 g/l) medium supplemented with 2 mM l -glutamine, 100 U/ml penicillin/streptomycin, and 10% fetal bovine serum. The Hela Kyoto BAC cell lines were kept under selection in geneticin (G-418, Thermo Fisher, 400 μg/ml).

Transfection, cDNAs, protein extraction, and immunoblotting

Transfections of cDNAs and siRNAs were performed using Lipofectamine 2000 (Life Technologies) following manufacturer's instructions. Experiments were performed 24 and 72 h after transfection of cDNAs or siRNA, respectively.

The cDNAs used were as follows: RPL23a-GFP (RG217630, OriGene) eGFP (Clonetech) GFP-GR50 (Lee et al, 2016 ) and Flag-VCP (Ritson et al, 2010 ) mCherry-PML was generated for this study GFP-TDP43 (Zhang et al, 2009 ) mCherry-VHL (Mateju et al, 2017 ) NLuc-GFP (Nollen et al, 2001 ) HA-Ubiquitin and Flag-Ubiquitin (den Engelsman et al, 2003 ).

To extract total proteins, cells were lysed in Laemmli sample buffer and homogenized by sonication. Prior to separation by SDS–PAGE, the protein samples were boiled for 3 min at 100°C and reduced with β-mercaptoethanol. Proteins were transferred onto nitrocellulose membranes and analyzed by Western blotting.

Nucleoli isolation

Nucleoli were isolated from HeLa cells as previously described (Andersen et al, 2002 ) with some variations. Briefly, 5 × 150 mm Petri dishes of confluent HeLa cells were washed three times with cold PBS. Cells were scraped and collected by centrifugation at 218 g and 4°C for 5 min. Cells were subsequently resuspended in 5 ml of cold Buffer A (10 mM Hepes pH 7.9, 10 mM KCl, 1.5 mM MgCl2, 0.5 mM DTT, EDTA-free protease inhibitors, Roche) and incubated 15 min on ice. Cells were then dounce homogenized 20 times using a tight pestle and centrifuged at 218 g and 4°C for 5 min. The supernatant was collected as cytoplasmic fraction, while the pellet, which contained nuclei, was resuspended with 3 ml of S1 solution (0.25 M sucrose, 10 mM MgCl2, EDTA-free protease inhibitors). Resuspended nuclei were layered over 3 ml of S2 solution (0.35 M sucrose, 0.5 mM MgCl2, EDTA-free protease inhibitors) and centrifuged at 1,430 g and 4°C for 5 min. The resulting nuclear pellet was resuspended with 3 ml of S2 solution and sonicated 6 × 10 s bursts at 20% amplitude (Branson Digital Sonifier 450-D). The sonicated sample was layered over 3 ml of S3 solution (0.88 M sucrose, 0.5 mM MgCl2, EDTA-free protease inhibitors) and centrifuged at 3,000 g and 4°C for 10 min. The supernatant was collected as nucleoplasmic fraction. The pellet, which contained nucleoli, was resuspended in 0.5 ml of S2 solution, centrifuged at 1,430 g and 4°C for 5 min, resuspended in 0.1 ml of S2 solution, and used to assess nucleoli purity and DRiP distribution by Western blotting.

Immunofluorescence on cultured cells and labeling of nascent peptides with OP-puro

Cells were grown on polylysine-coated glass coverslip. After washing with cold PBS, cells were fixed with 3.7% formaldehyde in PBS for 9 min at room temperature, followed by permeabilization with cold acetone for 5 min at −20°C. Alternatively, cells were fixed with ice-cold methanol for 10 min at −20°C. PBS containing 3% BSA and 0.1% Triton X-100 was used for blocking and incubation with primary and secondary antibodies.

Labeling of newly synthesized proteins was performed by incubating the cells with 25 μM O-propargyl-puromycin (OP-puro) for the indicated time points. OP-puro-labeled peptides were detected by click chemistry as previously described (Ganassi et al, 2016 ).

Amylo-glo staining of cultured cells

HeLa cells were seeded on non-coated glass coverslips. 24 h later, cells were either left untreated or subjected to stress as described in the main text. Cells were then washed with cold PBS, fixed with 3.7% formaldehyde in PBS for 9 min at room temperature, and permeabilized with PBS containing 0.2% Triton X-100. Cells were stained with Amylo-glo (1X) in 0.9% NaCl for 15 min at room temperature and washed for 5 min with 0.9% NaCl. Cells were immediately analyzed by confocal microscopy.

High content imaging-based assay

Images were obtained using a Leica SP2 AOBS system (Leica Microsystems) and a 63 × oil immersion lens. PML body number, DRiP foci number, and enrichment for DRiPs and polyUb proteins inside PML bodies were analyzed using the Scan R Analysis software (Olympus). First, PML bodies were segmented based on PML signal using edge detection algorithm. Following PML body segmentation, we measured the mean fluorescence intensity of the protein of interest in each detected PML body. Additionally, the mean fluorescence intensity of the protein was measured in an area surrounding the PML body. The relative enrichment of the protein in individual PML bodies was calculated as a ratio of the mean fluorescence intensity inside the PML body divided by the mean intensity in the region surrounding the PML body. The values were plotted as column graphs representing the fraction of PML bodies with enrichment > 1.5. Two-tailed t-test was performed to compare the enrichment values between two groups.

Live-cell imaging and Fluorescence recovery after photobleaching

Live-cell imaging was done using the DeltaVision imaging system and SoftWorx 4.1.2 and with the Leica SP8 system. FRAP measurements on PML-GFP HeLa Kyoto cells were performed using a spinning-disk confocal microscope (Olympus IX81), while FRAP measurements on GFP-PSMA7 HeLa Kyoto cells were performed using the Leica SP8 system.

For FRAP analysis on PML-GFP HeLa Kyoto cells, we used a 100× oil immersion objective. A region of approximately 2.02 × 2.02 μm was bleached for 60 ms using a laser intensity of 30% at 405 nm. Recovery was recorded for 120 time points after bleaching (600 s). For FRAP analysis on GFP-PSMA7 HeLa Kyoto cells, we used a 63× oil immersion objective. A region of approximately 2.2–2.5 × 2.2–2.5 μm was bleached for 1 s using a laser intensity of 30% at 405 nm. For FRAP analysis of untreated cells or in cells during the stress recovery in drug-free medium, a laser intensity of 100% for 5 s was used. Recovery was recorded for 600 time points after bleaching (600 s). Analysis of the recovery curves for both PML-GFP and GFP-PSMA7 were carried out with the FIJI/ImageJ.

The flow of the protein within the PML body or the GFP-PSMA7 enriched body was measured by quantifying the recovery of the bleached area at the cost of the unbleached region and using a custom written FIJI/ImageJ routine. The bleached region was corrected for general bleaching during image acquisition. We quantified the molecules that move from the unbleached region to the bleached region, leading to recovery of the bleached region.

Prior to FRAP analysis, we corrected the images for drift using the StackReg plug-in function of the FIJI software suite. The equation used for FRAP analysis is as follows ((Ibleach − Ibackground)/(Ibleach(t0) − Ibackground(to)))/((Itotal-Ibackground)/(Itotal(t0) − Ibackground(to))), where Itotal is the fluorescence intensity of the entire cellular structure, Ibleach represents the fluorescence intensity in the bleach area, and Ibackground the background of the camera offset. FRAP curves were averaged to obtain the mean and standard deviation. Fluorescent density analysis was performed using FIJI/ImageJ and selecting specific region of interest (ROI).

Colony formation assay

Colony formation assay was performed as previously described (Crowley et al, 2016 ). Briefly, 24 h after seeding, HeLa cells were either left untreated or subjected to proteotoxic stress as described in the main text. Cells were subsequently allowed to recover from stress and form colonies at 37°C, 5% CO2 for 10 days. Then, colonies were fixed with 100% methanol at room temperature for 20 min and stained with 0.5% crystal violet in 25% methanol for 5 min.

Quantification and statistical analysis

All statistical analyses were performed using one-way ANOVA, followed by Bonferroni–Holm post hoc test for comparisons between three or more groups or Student's t-test for comparisons between two groups using Daniel's XL Toolbox.

Materials and methods

Cultivation of Caco-2 cells

Human epithelial colorectal adenocarcinoma Caco-2 cells were obtained from the German Collection of Microorganisms and Cell Cultures (DSMZ, Cat. No. ACC 169) and routinely passaged in MEM medium (Gibco) with GlutaMAX and Earle’s salts, supplemented with 20% fetal calf serum (FCS, Biochrom), 1% Non-Essential Amino Acids (NEAA, Gibco), and 1% Sodium Pyruvate (Gibco) in a 5% CO2 humidified atmosphere at 37°C. Cells were cultured in T-75 (Sarstedt) flasks until 80–90% confluence and passaged at a sub-culture ratio of 1/2 to 1/10. For passaging, cells were washed once with DPBS (Dulbecco’s phosphate-buffered saline without CaCl2 and MgCl2, Gibco) to remove residual FCS and subsequently treated with 3 ml of 0.05% Trypsin-EDTA (Gibco) for 5 minutes at 37°C. The trypsinization process was stopped by addition of fresh cell culture medium to the flask and gentle mixing of the cells to give a homogenous single-cell suspension. Depending on the passaging ratio, a certain volume of this suspension was transferred to a new flask containing 10–13 ml of freshly supplemented cell culture medium and cells were given appropriate time to grow to confluence again.

Ethics statement

For preparation of the decellularized intestinal scaffold (SISmuc), porcine jejunal segments were explanted from six week-old piglets provided by the pig breeder Niedermayer from Dettelbach, Germany. The pigs were sacrificed after heparinization in accordance with the approval given by the government of Lower Franconia in Bavaria, Germany under the study number 55.2-2532-2-256.

Human primary biopsy samples were collected in collaboration with the surgical unit at the University Hospital Würzburg (PD Dr. med. Christian Jurowich, PD Dr. med. Florian Seyfried) from obese adults during routine stomach bypass surgery, and informed consent from patients was obtained beforehand. The use of human primary tissue was approved by the institutional ethics committee on human research of the Julius-Maximilians-University Würzburg under the number 182/10.

Generation of 3D tissue models and simulation of fluid dynamics

The SISmuc scaffold was generated by decellularization of porcine intestinal tissue via a standardized protocol published previously [35,36]. Briefly, after extensive washing with phosphate-buffered saline (PBS, Gibco), jejunal tissue was subjected to multiple rounds of decellularization with 4% sodium deoxycholate (Sigma Aldrich), alternating with washing steps in PBS. Subsequently, the tissue was digested with a DNase I solution (Roche) and sterilized using gamma radiation (BBFS Sterilisationsservice GmbH, Rommelhausen, Germany). SISmuc scaffolds were kept in this state in PBS at 4°C until usage. In order to generate 3D tissue models, the scaffolds were opened longitudinally, cut into 2 x 2 cm squares, and fixed between two metal rings (so-called cell crowns, Fraunhofer IGB, Stuttgart, Germany). This effectively created an apical and basolateral compartment with the former mucosal surface facing towards the apical side. The surface area of these scaffolds was previously estimated to be 3.1 cm 2 [36] and was seeded with 3 x 10 5 Caco-2 cells in 500 μl of the cell culture medium (MEM + 20% FCS + 1% NEAA + 1% Sodium Pyruvate), while the basolateral compartment was filled with 1.5 ml of the same medium. Tissue models were then routinely cultured either statically or dynamically on an orbital shaker (Celltron Infors HT) at 65 rpm in a 5% CO2 humidified atmosphere at 37°C for 21–28 days. Fresh medium was supplied every two days. 2D-Transwell inserts (polycarbonate, Ø 12 mm, 0.4/3.0 μm pore size, Corning), were similarly seeded with 3 x 10 5 Caco-2 cells and cultured statically with medium renewal every second day. Fluid dynamics to determine the rotary frequency of the shaker for dynamic culture conditions were simulated using COMSOL Multiphysics software (Comsol Multiphysics GmbH, Berlin, Germany) and were performed by Ivo Schwedhelm (Tissue Engineering and Regenerative Medicine, University Hospital Würzburg, Germany) as previously described [40].

Assessment of epithelial barrier integrity

Barrier integrity of the reconstructed mucosal tissue in cell crowns, as well as of confluent cell monolayers on 2D-Transwell inserts during either culture or infection with C. jejuni was assessed by determining the paracellular flux of fluorescein isothiocyanate-dextran (FITC-dextran). Specifically, 0.25 mg/ml of FITC-dextran (4 kDa, Sigma Aldrich) was resuspended in 500 μl of cell culture medium containing 20% FCS, 1% NEAA, and 1% Sodium Pyruvate, and applied to the apical compartment of tissue models or 2D-Transwells. Prior to that, 1.5 ml of the same culture medium without FITC-dextran was used to fill the basolateral compartment. From both the initial FITC-dextran solution as well as the fresh basolateral cell culture medium, three 100 μl samples were taken as a positive and negative control, respectively, and pipetted into separate wells of a black 96-well plate. Cell culture models were incubated with the FITC-dextran solution in their apical compartments at 37°C for 30 minutes. Afterwards, three 100 μl samples were taken from each basolateral compartment and measured for fluorescence intensity using an Infinite 200 PRO plate reader (TECAN). Autofluorescence of the pure cell culture medium was subtracted from total fluorescence intensities, and FITC-dextran permeability values for each cell culture model were calculated relative to the fluorescence intensity of the initial FITC-dextran solution. Barrier integrity was assessed routinely every seven days during culture. During infection experiments, permeability was measured every 24 hours for the same infected crown as well as for non-infected control cell culture models.

Histological and immunofluorescence staining and imaging

For histological staining, human small intestinal biopsy samples and 3D tissue models were fixed with 2% PFA (paraformaldehyde, Roth) for at least 1 hour at room temperature (RT), processed for paraffin embedding using a Leica ASP200S tissue processor (Leica Biosystems), and sectioned with 5 μm thickness using a Leica RM2255 microtome (Leica Biosystems). After deparaffinization and rehydration, sections were stained with Hematoxylin and Eosin (H&E adapted from [94]). After the staining process, light microscopy images were obtained with a Leica DM4000 B microscope (Leica Microsystems).

For immunofluorescence staining, tissue was processed as described above until the deparaffinization and rehydration of the tissue sections was completed. Subsequently, antigens were retrieved by incubation in 10 mM sodium citrate, pH 6 at 90°C for 20 minutes. When staining for C. jejuni, an additional enzymatic antigen retrieval step was applied by incubation with Proteinase K (Dako) for 12 minutes at RT. Sections were then permeabilized in DPBS with 0.5% Triton X-100 (Roth) for 30 minutes at RT and subsequently blocked with 1% bovine serum albumin (BSA, Roth) in PBS for 30 minutes at RT. Next, sections were incubated overnight at 4°C in a humidity chamber with primary antibody solution (DCS labline), followed by washing 3 x 15 minutes with PBS + 0.05% Tween 20 (Roth). Subsequently, tissue sections were incubated with the appropriate secondary antibody solution containing DAPI (4′,6-diamidino-2-phenylindole, Sigma Aldrich) for 4 hours at RT in the dark. After three washing steps with PBS + 0.05% Tween 20 for 15 minutes each at RT, samples were mounted in Mowiol (Sigma Aldrich) and imaged with a laser scanning Leica TCS SP5 II confocal microscope (Leica Microsystems) using z-stack scanning mode. All primary antibodies were diluted 1:100 and included monoclonal mouse anti-E-cadherin (BD Biosciences, 610181), monoclonal mouse anti-occludin (ThermoFisher Scientific, OC3F10), polyclonal rabbit anti-Campylobacter jejuni (GeneTex, GTX40882), and polyclonal rabbit anti-cytokeratin-18 (abcam, 1924–1). All secondary antibodies were diluted 1:400 and included goat anti-mouse IgG AlexaFluor555-conjugate (ThermoFisher Scientific, A21424), goat anti-mouse IgG AlexaFluor647-conjugate (ThermoFisher Scientific, A21236), donkey anti-rabbit IgG AlexaFluor594-conjugate (ThermoFisher Scientific, A21207), goat anti-rabbit IgG AlexaFluor647-conjugate (ThermoFisher Scientific, A21246).

Determination of cell height

For Caco-2 cells grown on 2D-Transwell inserts and SISmuc (static and dynamic culture), as well as for native human intestinal tissues, the mean height of cells was determined as follows. Tissues were fixed with 2% PFA and processed according to the immunofluorescence staining procedure described above. Ten confocal microscopy images were taken for five different Caco-2 cell-based 2D-Transwell inserts and tissue models (static and dynamic culture), respectively. In addition, ten images of similarly processed small intestinal patient samples (n = 5) were used to measure the cell height of every second cell in native tissue. Measurements were made using the program ImageJ. Altogether, 269 cells were measured for Caco-2 cells cultured on 2D-Transwell inserts, 271 cells for statically cultured Caco-2 models, 277 cells for dynamic tissue models, and 199 cells for native human intestine.

Determination of cell numbers on the Caco-2 3D tissue models

In order to assess the final number of epithelial cells present on a fully developed tissue model (21 days of culture), two methods were employed. First, tissue models were subjected to 30 minutes treatment with 0.05% Trypsin-EDTA at 37°C, cells were detached by scratching with a pipette tip, and then finally counted using a Neubauer counting chamber. The cell numbers for 24 statically cultured and 12 dynamically cultured tissue models were counted (S1 and S3 Tables), respectively. Second, the amount of DNA present in a tissue model was measured using the Quant-iT PicoGreen dsDNA assay kit (ThermoFisher Scientific) according to the manufacturer’s instructions. Briefly, DNA was isolated from 300,000 and 600,000 Caco-2 cells and quantified relative to a previously determined standard curve in order to correlate the DNA amount to a defined number of Caco-2 cells. DNA was also isolated from three statically grown Caco-2 tissue models and three unseeded SISmuc scaffolds and equally quantified. Based on these nucleic acid amounts, cell numbers in the tissue models (minus the residual DNA content of empty scaffolds), were calculated (S2 Table).

Bacterial strains, oligonucleotides, and plasmids

C. jejuni strains used in this study are listed in S4 Table. DNA oligonucleotides used for cloning of C. jejuni mutant strains are listed in S5 Table. Plasmids are listed in S6 Table.

C. jejuni standard growth conditions

All C. jejuni strains were routinely grown at 37°C on Mueller-Hinton (MH, Becton Dickinson) agar plates supplemented with 10 μg/ml vancomycin for 1–2 passages in a HERAcell 150i incubator (ThermoFisher Scientific) in a microaerobic environment (10% CO2, 5% O2, 85% N2). Agar plates were further supplemented with marker-selective antibiotics (20 μg/ml chloramphenicol, 250 μg/ml hygromycin, 50 μg/ml kanamycin) for selection of transformed clones. Bacteria were then transferred to Brucella Broth (BB, Becton Dickinson) liquid cultures in T25 flasks (Corning) by inoculation from plate to a final OD600 of 0.005 and grown under agitation at 140 rpm and 37°C.

Motility assay

Liquid cultures of C. jejuni strains in BB media containing 10 μg/ml vancomycin were grown under agitation to mid-log phase (OD600 0.4) at 37°C in a microaerobic environment. For each strain, 0.5 μl of bacterial culture was inoculated into a motility soft-agar plate (BB broth + 0.4% Difco agar) poured one day prior to the experiment. Plates were incubated right-side-up until halo formation could be observed (approximately 12–20 hrs post inoculation). For each inoculation, halo radius was measured three times and averaged to give the mean swimming distance for each strain on each plate. Each strain was inoculated in technical triplicates per experiment and motility assays were performed in three independent biological replicates. The average halo radius for each strain was used to compare motility between C. jejuni wild-type and mutant strains.

Construction of C. jejuni deletion mutant strains

Deletion mutants of C. jejuni used in this study were constructed by double-crossover homologous recombination with antibiotic resistance cassettes to remove most of the coding sequence in the genomic locus thereby disrupting the respective genes. Resistance cassettes used for cloning were either aphA-3 (Kan R ) [95], aph(7”) (Hyg R ) [96], or cat.coli (Cm R ) [97]. Non-polar resistance cassettes were amplified with primers HPK1/HPK2 from plasmid pGG1 [58] for Kan R , with primers CSO-1678/CSO-1679 from plasmid pAC1H [96] for Hyg R , or primers CSO-0613/0614 from C. jejuni strain CSS-0643 [58] for Cm R . Overlap PCR products carried these resistance cassettes flanked by

500 bp of homologous sequence up- and downstream of the gene to be deleted. As an example, deletion of kpsMT in C. jejuni strain NCTC11168 (CSS-0032) will be described in detail. First,

500 bp upstream of the kpsM (Cj1448c) start codon using CSO-2009 and CSO-2008 and

500 bp downstream of the kpsM stop codon using CSO-2011 and CSO-2010 were amplified from genomic DNA of NCTC11168 wild-type strain. As the start codon of the downstream gene kpsT (Cj1447c) overlaps with the stop codon of kpsM, the mutant created here likely resulted in the inactivation of kpsT as well. The kanamycin resistance cassette (aphA-3) was amplified from pGG1 using HPK1 and HPK2. To fuse the up- and downstream region of kpsM with the aphA-3 resistance cassette, the antisense oligonucleotide of the kpsM upstream region (CSO-2008) contained 22 bp of overlap with the sense oligonucleotide used to amplify the aphA-3 resistance cassette (HPK1). Likewise, the sense oligonucleotide of the kpsM downstream region (CSO-2011) contained 26 bp overlap with the antisense oligonucleotide to amplify the aphA-3 resistance cassette (HPK2). In a final 100 μl Phusion polymerase PCR reaction, the purified (Macherey-Nagel NucleoSpin PCR cleanup kit) up- and downstream regions of kpsM were added together with the aphA-3 resistance cassette in a ratio of 50:50:90 ng and amplified using CSO-2009 and CSO-2011 (final concentration of 1 μM). The program for the overlap PCR was as follows: 1 cycle of [98°C, 1 min 61°C, 1 min 72°C, 10 min 98°C, 1 min], 40 cycles of [98°C, 15 s 57°C, 30 s 72°C, 1 min], followed by 72°C for 10 min. Overlap PCR products were verified for their size by agarose gel electrophoresis and after purification subsequently transformed into the recipient C. jejuni strain by electroporation (see protocol described below). After verification of the resulting clones via colony PCR with CSO-2008 and HPK2, a positive clone was picked for the final kpsMT deletion strain (CSS-6198 NCTC11168 ΔkpsMT). Deletion mutants for flaA (CSS-1512) [58], csrA (CSS-0643) [58], cas9 (CSS-3836) [78], and ptmG (CSS-2966) in C. jejuni strain NCTC11168, as well as for flaA (CSS-2380), kpsMT (CSS-6200), and csrA (CSS-6202) in strain 81–176 (CSS-0063) were constructed using an analogous approach and oligonucleotides are listed in S5 Table.

The sRNA locus CJnc180/190 was deleted in the same way as described with the exception of using a resistance cassette that included a promoter and a terminator sequence. This resistance cassette was amplified using JVO-5068 and HPK2term from pGG1 and annealed together with the purified up- and downstream fragments of CJnc180/190. The up- and downstream regions of CJnc180/190 were amplified using CSO-0247/CSO-0248 and CSO-0249/CSO-0250, respectively, from wild-type genomic DNA of strain NCTC11168. CSO-0248 and CSO-0249 contained overlapping regions to the polar kanamycin resistance cassette with a promoter and terminator region from the H. pylori sRNA RepG [98]. The upstream region and downstream region as well as cassette PCR amplicons were then annealed and the entire product was amplified with CSO-0247 and CSO-0250 and electroporated into the C. jejuni NCTC11168 wild-type strain. Kanamycin-resistant colonies were validated via colony PCR using CSO-0246 and HPK1, resulting in CJnc180/190::aphA-3 (CSS-1157).

Construction of C. jejuni complementation and overexpression strains

In order to complement the deletion of a gene or create an overexpression construct, a wild-type copy of the gene of interest was inserted into the rdxA (Cj1066) locus, which is frequently used for complementation in C. jejuni [99]. To achieve this, complementation/overexpression constructs were first generated in plasmids that contained approximately 500 bp up- and downstream sequences of the insertion site in the rdxA gene, flanking Cm R or Kan R resistance cassettes that each contained a promoter and terminator, or were generated by overlap PCR.

As an example for the plasmid-based approach, the construction of the ptmG complementation and overexpression in strain NCTC11168 is described in detail. The coding region of ptmG including approximately 100 nt up- and downstream of the start and stop codon, respectively, was amplified from genomic DNA of C. jejuni strain NCTC11168 using CSO-2928 and CSO-2929 and digested with XmaI and PstI. The backbone for the ptmG complementation/overexpression vector was amplified by inverse PCR with CSO-0762 and CSO-0493 from pST1.1 [78] and likewise digested (XmaI/PstI). The plasmid backbone and the insert were ligated and transformed into E. coli TOP10. The clones were verified by colony PCR using CSO-0023 and CSO-2929 and sequenced using CSO-0023 (Macrogen), resulting in pSSv63.1. A purified complementation construct, amplified from pSSv63.1 with CSO-2276 and CSO-2277, was then transformed into C. jejuni strain NCTC11168 ΔptmG (CSS-2966) via electroporation for complementation or into NCTC11168 WT (CSS-0032) for overexpression of ptmG. The final complementation/overexpression clones were verified by colony PCR using CSO-0023 and CSO-0349 and sequencing using CSO-0023, resulting in the complementation strain C ptmG (CSS-2978 ptmG::aph(7”) rdxA::aphA-3-ptmG) and overexpression strain OE ptmG (CSS-2980 rdxA::aphA-3-ptmG), respectively. An analogous approach was carried out for C CJnc180/190 (CSS-1158 CJnc180/190::aphA-3, rdxA::cat.coli-CJnc180/190) in their respective deletion mutant strains.

Generation of complementation constructs by overlap PCR (i.e., rxdA::cat.coli-flaA, rdxA::cat.coli-kpsMT, and rdxA::aphA-3-csrA) was performed as follows, using NCTC11168 flaA as an example. A fragment encoding the rdxA upstream region (approximately 500 bp) fused to the cat.coli Cm R resistance cassette (with promoter and terminator) was amplified by PCR from pGD34.7 using CSO-2276/CSO-0573, and the rdxA downstream region was amplified with CSO-0347/CSO-2277 from NCTC11168 wild-type genomic DNA. The flaA gene, with its native promoter [55], was amplified from NCTC11168 wild-type genomic DNA using CSO-4744/CSO-4745, such that the 5’-end of the amplicon had complementarity to CSO-0573, and the 3’-end of the amplicon overlapped with CSO-0347. The rdxA_up-cat.coli, flaA insert, and rdxA_down fragments were then annealed, and the entire product was amplified with CSO-2276 and CSO-2277. The resulting PCR product was electroporated into C. jejuni NCTC11168 ΔflaA, and insertion mutants were validated by colony PCR with CSO-0643/CSO-0349 and sequencing with CSO-0643/CSO-4474/CSO-3270. The same approach was used for complementation of ΔkpsMT strains. For complementation of ΔcsrA, a similar approach was used, except the upstream fragment encoded rdxA_up-aphA-3. This fragment was amplified from pST1.1 with CSO-2276/CSO-0762. The csrA insert for both strains included a putative promoter encoded within the upstream truB locus, identified by differential RNA-seq [55]. Complemented csrA strains were validated by colony PCR with CSO-0023/CSO-3270 and sequencing with CSO-0023.

Transformation of C. jejuni by electroporation

C. jejuni NCTC11168 or 81–176 wildtype or appropriate deletion mutants were streaked onto MH agar plates with the suitable antibiotics from cryostocks. After one passage, bacterial cells were harvested with a cotton swab and resuspended in cold electroporation solution (272 mM sucrose, 15% (w/v) glycerol). Bacteria were harvested by centrifugation at 4°C and 6,500 x g for 5 minutes, and then resuspended in the same solution. After two additional washing steps, the final pellet was resuspended in an appropriate small volume of electroporation solution, depending on the size of the pellet. Next, 50 μl of this cell suspension was mixed with 200–400 ng of purified PCR product (not exceeding 4 μl in total) and electroporated (Biorad Genepulser) in a 1 mm gap cuvette (PEQLAB) at 2.5 kV, 200 Ω, and 25 μF. By adding 200 μl prewarmed Brucella Broth, cells were then transferred onto a non-selective MH agar plate and recovered overnight at 37°C in a microaerobic environment. The next day, bacterial cells were harvested with a cotton swab, streaked onto an appropriate selective MH agar plate, and incubated at 37°C microaerobically until colonies were observed (typically 2–4 days). Clones were verified by colony PCR and sequencing, cryostocks were frozen in 25% glycerol in BB medium, and stored at -80°C.

Infection of the 3D tissue model with C. jejuni

Caco-2 cell-based tissue models were cultured either statically or dynamically for 21 days before they were used for infection experiments with C. jejuni. Independent of pre-culture of the tissue models, infection with C. jejuni was always conducted under static conditions without additional mechanical stimulation. Bacteria were grown as described above, harvested from liquid culture in mid-log phase (OD600 0.4), and resuspended in fresh cell culture medium containing 20% FCS, 1% NEAA, and 1% Sodium Pyruvate to achieve a multiplicity of infection (MOI) of 20. From this bacterial cell suspension, serial dilutions were plated onto MH agar plates and incubated at 37°C under microaerobic conditions to determine the input amount of colony forming units (CFUs). 500 μl of these bacterial suspensions were used to apically infect the tissue models and co-incubation was carried out in a 5% CO2 humidified atmosphere at 37°C. For transmigration experiments, 100 μl samples from the basolateral compartment of infected tissue models were taken at indicated time points post infection (10 min to 8 hrs p.i.) and serial dilutions were plated on MH agar plates to determine the number of transmigrated bacteria. To isolate bacteria adherent to and internalized into the tissue models, infection was stopped at indicated time points post infection (4–120 hrs). For experiments involving longer time points (24–120 hrs), spent cell culture medium was exchanged daily for fresh cell culture medium supplemented with 20% FCS, 1% NEAA, and 1% Sodium Pyruvate in the apical and the basolateral compartment of the tissue models. To harvest cell-associated bacteria, cell crowns were washed three times with DPBS to remove all non-adherent bacterial cells. Subsequently, two tissue pieces per crown were collected using a tissue punch (Ø 5 mm, Kai Medical) and transferred to an Eppendorf tube with 500 μl of 0.1% saponin in DPBS. The tissue pieces were incubated for 10 minutes at 37°C under agitation to isolate bacteria from host cells. Serial dilutions were then plated on MH agar plates, colonies were counted, and CFU numbers were calculated as a percentage of input CFUs for each strain. In order to specifically isolate host cell-internalized C. jejuni, fresh cell culture medium with 20% FCS, 1% NEAA, and 1% Sodium Pyruvate containing 200 μg/ml gentamicin was supplied to the basolateral and the apical compartment of the tissue models to kill extracellular bacteria for 2 hours at 37°C. This concentration of gentamicin was determined by subjecting both C. jejuni 81–176 and NCTC11168 to varying concentrations of gentamicin (10–400 μg/ml) for different periods of time (30–240 min). Plating for CFUs was used as a readout for bacteria surviving the antibiotic treatment. After treatment with 200 μg/ml for 2 hrs, no colonies could be recovered for either of the wild-type strains. In addition, CFU plating of the supernatant for each C. jejuni strain after this gentamicin treatment ensured that no extracellular bacteria had survived. For experiments involving intracellular survival of C. jejuni in the tissue model, the initial treatment with 200 μg/ml of gentamicin for 2 hrs was followed by replacement of the medium in the basolateral and the apical compartment for fresh cell culture medium containing 20% FCS, 1% NEAA, and 1% Sodium Pyruvate supplemented with 10 μg/ml gentamicin. Again, CFU plating of the supernatant ensured that no extracellular bacteria could be recovered after this treatment. After gentamicin treatment, tissue models were washed three times with DPBS before CFU numbers were determined as described above.

Infection of 2D cell culture models (2D-monolayer and 2D-Transwell) with C. jejuni

In general, infection experiments were carried out as described for 3D tissue models with a few modifications. For 2D-monolayer infections, Caco-2 cells were seeded into 6-well plates two days prior to the infection experiment in order to achieve a confluent cell monolayer. C. jejuni was grown in liquid culture to mid-log phase, resuspended in cell culture medium and used for infection of the epithelial monolayer at an MOI of 20. After infection, cells were washed three times with DPBS, lysed with 0.1% saponin in 1 ml DPBS, and the resulting cell suspension was plated in serial dilutions on MH agar plates. For specific recovery of intracellular bacteria, cells were treated with 200 μg/ml gentamicin for 2 hrs at 37°C and CFUs were determined as described. In addition, CFU plating of the supernatant of gentamicin-treated wells ensured that all extracellular bacteria were killed during the antibiotic treatment. For 2D-Transwell infections, Caco-2 cells were grown on polycarbonate 2D-Transwell inserts (Corning, 12 mm, 3.0 μm) for 21 days in a 5% CO2 humidified atmosphere at 37°C. Isolation and enumeration of CFU numbers of colonizing or transmigrated bacteria was carried out the same way as described above for the 3D tissue models.

Growth curve analysis in cell culture medium and infection supernatants

To analyze the growth behavior of C. jejuni WT strains in tissue culture medium, bacteria were first routinely grown on MH agar plates and subsequently in BB liquid cultures overnight as described above. The next morning, bacterial cells were washed once with PBS and inoculated to a final OD600 of 0.05 into 50 ml cell culture medium (MEM + 20% FCS, 1% NEAA, 1% Sodium Pyruvate) and grown under agitation at 140 rpm in a HERAcell 150i incubator (ThermoFisher Scientific) in a microaerobic environment (10% CO2, 5% O2, 85% N2). Samples were removed at appropriate time-points post-inoculation for plating on MH agar to determine CFUs/ml. To measure growth of C. jejuni wild-type strains in the supernatant above 3D tissue models and 2D-Transwells, infection experiments with C. jejuni strains NCTC11168 and 81–176 were carried out as described above with the exception that the cell culture medium was not exchanged daily. At the appropriate time points post-inoculation, 10 μl samples were taken from the supernatant for plating on MH agar plates to ascertain CFUs/ml.

Kim, Y. G., Cha, J. & Chandrasegaran, S. Hybrid restriction enzymes: zinc finger fusions to Fok I cleavage domain. Proc. Natl Acad. Sci. USA 93, 1156–1160 (1996).

Wolfe, S. A., Nekludova, L. & Pabo, C. O. DNA recognition by Cys2His2 zinc finger proteins. Annu. Rev. Biophys. Biomol. Struct. 29, 183–212 (2000).

Bibikova, M., Beumer, K., Trautman, J. K. & Carroll, D. Enhancing gene targeting with designed zinc finger nucleases. Science 300, 764 (2003).

Miller, J. C. et al. An improved zinc-finger nuclease architecture for highly specific genome editing. Nat. Biotechnol. 25, 778–785 (2007).

Porteus, M. H. & Baltimore, D. Chimeric nucleases stimulate gene targeting in human cells. Science 300, 763 (2003).

Boch, J. et al. Breaking the code of DNA binding specificity of TAL-type III effectors. Science 326, 1509–1512 (2009).

Moscou, M. J. & Bogdanove, A. J. A simple cipher governs DNA recognition by TAL effectors. Science 326, 1501 (2009).

Christian, M. et al. Targeting DNA double-strand breaks with TAL effector nucleases. Genetics 186, 757–761 (2010).

Miller, J. C. et al. A TALE nuclease architecture for efficient genome editing. Nat. Biotechnol. 29, 143–148 (2011).

Mahfouz, M. M. et al. De novo-engineered transcription activator-like effector (TALE) hybrid nuclease with novel DNA binding specificity creates double-strand breaks. Proc. Natl Acad. Sci. USA 108, 2623–2628 (2011).

Li, T. et al. TAL nucleases (TALNs): hybrid proteins composed of TAL effectors and FokI DNA-cleavage domain. Nucleic Acids Res. 39, 359–372 (2011).

Jinek, M. et al. A programmable dual-RNA-guided DNA endonuclease in adaptive bacterial immunity. Science 337, 816–821 (2012).

Jansen, R., Embden, J. D., Gaastra, W. & Schouls, L. M. Identification of genes that are associated with DNA repeats in prokaryotes. Mol. Microbiol. 43, 1565–1575 (2002).

Gasiunas, G., Barrangou, R., Horvath, P. & Siksnys, V. Cas9-crRNA ribonucleoprotein complex mediates specific DNA cleavage for adaptive immunity in bacteria. Proc. Natl Acad. Sci. USA 109, E2579–E2586 (2012).

Cong, L. et al. Multiplex genome engineering using CRISPR/Cas systems. Science 339, 819–823 (2013).

Jeggo, P. A. DNA breakage and repair. in Advances in Genetics Vol. 38 (eds Hall, J. et al.) 185–218 (Academic Press, 1998).

Rouet, P., Smih, F. & Jasin, M. Introduction of double-strand breaks into the genome of mouse cells by expression of a rare-cutting endonuclease. Mol. Cell. Biol. 14, 8096–8106 (1994).

Lukacsovich, T., Yang, D. & Waldman, A. S. Repair of a specific double-strand break generated within a mammalian chromosome by yeast endonuclease I-Scel. Nucleic Acids Res. 22, 5649–5657 (1994).

Yeh, C. D., Richardson, C. D. & Corn, J. E. Advances in genome editing through control of DNA repair pathways. Nat. Cell Biol. 21, 1468–1478 (2019).

Haapaniemi, E., Botla, S., Persson, J., Schmierer, B. & Taipale, J. CRISPR–Cas9 genome editing induces a p53-mediated DNA damage response. Nat. Med. 24, 927–930 (2018).

Ihry, R. J. et al. p53 inhibits CRISPR–Cas9 engineering in human pluripotent stem cells. Nat. Med. 24, 939–946 (2018).

Shen, M. W. et al. Predictable and precise template-free CRISPR editing of pathogenic variants. Nature 563, 646–651 (2018).

van Overbeek, M. et al. DNA repair profiling reveals nonrandom outcomes at Cas9-mediated breaks. Mol. Cell 63, 633–646 (2016).

Koike-Yusa, H., Li, Y., Tan, E. P., Velasco-Herrera, M. D. C. & Yusa, K. Genome-wide recessive genetic screening in mammalian cells with a lentiviral CRISPR-guide RNA library. Nat. Biotechnol. 32, 267–273 (2014).

Kosicki, M., Tomberg, K. & Bradley, A. Repair of double-strand breaks induced by CRISPR–Cas9 leads to large deletions and complex rearrangements. Nat. Biotechnol. 36, 765–771 (2018).

Adikusuma, F. et al. Large deletions induced by Cas9 cleavage. Nature 560, E8–E9 (2018).

Jasin, M. & Rothstein, R. Repair of strand breaks by homologous recombination. Cold Spring Harb. Perspect. Biol. 5, a012740 (2013).

Paquet, D. et al. Efficient introduction of specific homozygous and heterozygous mutations using CRISPR/Cas9. Nature 533, 125–129 (2016).

Rees, H. A., Yeh, W.-H. & Liu, D. R. Development of hRad51–Cas9 nickase fusions that mediate HDR without double-stranded breaks. Nat. Commun. 10, 2212 (2019).

Richardson, C. D., Ray, G. J., Dewitt, M. A., Curie, G. L. & Corn, J. E. Enhancing homology-directed genome editing by catalytically active and inactive CRISPR-Cas9 using asymmetric donor DNA. Nat. Biotechnol. 34, 339–344 (2016).

Lin, S., Staahl, B. T., Alla, R. K. & Doudna, J. A. Enhanced homology-directed human genome engineering by controlled timing of CRISPR/Cas9 delivery. eLife 3, e04766 (2014).

Landrum, M. J. et al. ClinVar: public archive of relationships among sequence variation and human phenotype. Nucleic Acids Res. 42, D980–D985 (2013).

Landrum, M. J. et al. ClinVar: public archive of interpretations of clinically relevant variants. Nucleic Acids Res. 44, D862–D868 (2015).

Stenson, P. D. et al. The Human Gene Mutation Database: towards a comprehensive repository of inherited mutation data for medical research, genetic diagnosis and next-generation sequencing studies. Hum. Genet. 136, 665–677 (2017).

Komor, A. C., Kim, Y. B., Packer, M. S., Zuris, J. A. & Liu, D. R. Programmable editing of a target base in genomic DNA without double-stranded DNA cleavage. Nature 533, 420–424 (2016).

Gaudelli, N. M. et al. Programmable base editing of A•T to G•C in genomic DNA without DNA cleavage. Nature 551, 464–471 (2017).

Mok, B. Y. et al. A bacterial cytidine deaminase toxin enables CRISPR-free mitochondrial base editing. Nature 583, 631–637 (2020).

Nishimasu, H. et al. Crystal structure of Cas9 in complex with guide RNA and target DNA. Cell 156, 935–949 (2014).

Huang, T. P. et al. Circularly permuted and PAM-modified Cas9 variants broaden the targeting scope of base editors. Nat. Biotechnol. 37, 626–631 (2019).

Jiang, W. et al. BE-PLUS: a new base editing tool with broadened editing window and enhanced fidelity. Cell Res. 28, 855–861 (2018).

Ma, Y. et al. Targeted AID-mediated mutagenesis (TAM) enables efficient genomic diversification in mammalian cells. Nat. Methods 13, 1029–1035 (2016).

Hess, G. T. et al. Directed evolution using dCas9-targeted somatic hypermutation in mammalian cells. Nat. Methods 13, 1036–1042 (2016).

Liu, L. D. et al. Intrinsic nucleotide preference of diversifying base editors guides antibody ex vivo affinity maturation. Cell Rep. 25, 884–892.3 (2018).

Wang, Y., Zhou, L., Liu, N. & Yao, S. BE-PIGS: a base-editing tool with deaminases inlaid into Cas9 PI domain significantly expanded the editing scope. Signal Transduct. Target. Ther. 4, 36 (2019).

Kim, Y. B. et al. Increasing the genome-targeting scope and precision of base editing with engineered Cas9-cytidine deaminase fusions. Nat. Biotechnol. 35, 371–376 (2017).

Grünewald, J. et al. Transcriptome-wide off-target RNA editing induced by CRISPR-guided DNA base editors. Nature 569, 433–437 (2019).

Tan, J., Zhang, F., Karcher, D. & Bock, R. Engineering of high-precision base editors for site-specific single nucleotide replacement. Nat. Commun. 10, 439 (2019).

Pluciennik, A. et al. PCNA function in the activation and strand direction of MutLα endonuclease in mismatch repair. Proc. Natl Acad. Sci. USA 107, 16066–16071 (2010).

Heller, R. C. & Marians, K. J. Replisome assembly and the direct restart of stalled replication forks. Nat. Rev. Mol. Cell Biol. 7, 932–943 (2006).

Kurt, I. C. et al. CRISPR C-to-G base editors for inducing targeted DNA transversions in human cells. Nat. Biotechnol. (2020).

Zhao, D. et al. Glycosylase base editors enable C-to-A and C-to-G base changes. Nat. Biotechnol. (2020).

Kim, H. S., Jeong, Y. K., Hur, J. K., Kim, J.-S. & Bae, S. Adenine base editors catalyze cytosine conversions in human cells. Nat. Biotechnol. 37, 1145–1148 (2019).

Arbab, M. et al. Determinants of base editing outcomes from target library analysis and machine learning. Cell 182, 463–480.e30 (2020).

Nishida, K. et al. Targeted nucleotide editing using hybrid prokaryotic and vertebrate adaptive immune systems. Science 353, aaf8729 (2016).

Komor, A. C. et al. Improved base excision repair inhibition and bacteriophage Mu Gam protein yields C:G-to-T:A base editors with higher efficiency and product purity. Sci. Adv. 3, eaao4774 (2017).

Landrum, M. J. et al. ClinVar: improvements to accessing data. Nucleic Acids Res. 48(D1), D835–D844 (2020).

Koblan, L. W. et al. Improving cytidine and adenine base editors by expression optimization and ancestral reconstruction. Nat. Biotechnol. 36, 843–846 (2018).

Zafra, M. P. et al. Optimized base editors enable efficient editing in cells, organoids and mice. Nat. Biotechnol. 36, 888–893 (2018).

Wang, L. et al. Enhanced base editing by co-expression of free uracil DNA glycosylase inhibitor. Cell Res. 27, 1289–1292 (2017).

Zhang, X. et al. Increasing the efficiency and targeting range of cytidine base editors through fusion of a single-stranded DNA-binding protein domain. Nat. Cell Biol. 22, 740–750 (2020).

Cheng, T.-L. et al. Expanding C–T base editing toolkit with diversified cytidine deaminases. Nat. Commun. 10, 3612 (2019).

Li, C. et al. Targeted, random mutagenesis of plant genes with dual cytosine and adenine base editors. Nat. Biotechnol. 38, 875–882 (2020).

Sakata, R. C. et al. Base editors for simultaneous introduction of C-to-T and A-to-G mutations. Nat. Biotechnol. 38, 865–869 (2020).

Grünewald, J. et al. A dual-deaminase CRISPR base editor enables concurrent adenine and cytosine editing. Nat. Biotechnol. 38, 861–864 (2020).

Zhang, X. et al. Dual base editor catalyzes both cytosine and adenine base conversions in human cells. Nat. Biotechnol. 38, 856–860 (2020).

Levy, J. M. et al. Cytosine and adenine base editing of the brain, liver, retina, heart and skeletal muscle of mice via adeno-associated viruses. Nat. Biomed. Eng. 4, 97–110 (2020).

Villiger, L. et al. Treatment of a metabolic liver disease by in vivo genome base editing in adult mice. Nat. Med. 24, 1519–1525 (2018).

Yeh, W.-H. et al. In vivo base editing restores sensory transduction and transiently improves auditory function in a mouse model of recessive deafness. Sci. Transl. Med. 12, eaay9101 (2020).

Ryu, S.-M. et al. Adenine base editing in mouse embryos and an adult mouse model of Duchenne muscular dystrophy. Nat. Biotechnol. 36, 536–539 (2018).

Yang, L. et al. Amelioration of an inherited metabolic liver disease through creation of a de novo start codon by cytidine base editing. Mol. Ther. 28, 1673–1683 (2020).

Kleinstiver, B. P. et al. Engineered CRISPR-Cas9 nucleases with altered PAM specificities. Nature 523, 481–485 (2015).

Kleinstiver, B. P. et al. Broadening the targeting range of Staphylococcus aureus CRISPR-Cas9 by modifying PAM recognition. Nat. Biotechnol. 33, 1293–1298 (2015).

Kleinstiver, B. P. et al. High-fidelity CRISPR–Cas9 nucleases with no detectable genome-wide off-target effects. Nature 529, 490–495 (2016).

Nishimasu, H. et al. Engineered CRISPR-Cas9 nuclease with expanded targeting space. Science 361, 1259–1262 (2018).

Hu, J. H. et al. Evolved Cas9 variants with broad PAM compatibility and high DNA specificity. Nature 556, 57–63 (2018).

Ran, F. A. et al. In vivo genome editing using Staphylococcus aureus Cas9. Nature 520, 186–191 (2015).

Walton, R. T., Christie, K. A., Whittaker, M. N. & Kleinstiver, B. P. Unconstrained genome targeting with near-PAMless engineered CRISPR-Cas9 variants. Science 368, 290 (2020).

Miller, S. M. et al. Continuous evolution of SpCas9 variants compatible with non-G PAMs. Nat. Biotechnol. 38, 471–481 (2020).

Toth, E. et al. Improved LbCas12a variants with altered PAM specificities further broaden the genome targeting range of Cas12a nucleases. Nucleic Acids Res. 48, 3722–3733 (2020).

Chatterjee, P., Jakimo, N. & Jacobson, J. M. Minimal PAM specificity of a highly similar SpCas9 ortholog. Sci. Adv. 4, eaau0766 (2018).

Chatterjee, P. et al. An engineered ScCas9 with broad PAM range and high specificity and activity. Nat. Biotechnol. 38, 1154–1158 (2020).

Chatterjee, P. et al. A Cas9 with PAM recognition for adenine dinucleotides. Nat. Commun. 11, 2474 (2020).

Cox, D. B. T., Platt, R. J. & Zhang, F. Therapeutic genome editing: prospects and challenges. Nat. Med. 21, 121–131 (2015).

Anzalone, A. V., Koblan, L. W. & Liu, D. R. Genome editing with CRISPR–Cas nucleases, base editors, transposases and prime editors. Nat. Biotechnol. 38, 824–844 (2020).

Yang, L. et al. Increasing targeting scope of adenosine base editors in mouse and rat embryos through fusion of TadA deaminase with Cas9 variants. Protein Cell 9, 814–819 (2019).

Kleinstiver, B. P. et al. Engineered CRISPR–Cas12a variants with increased activities and improved targeting ranges for gene, epigenetic and base editing. Nat. Biotechnol. 37, 276–282 (2019).

Zhang, Y. et al. Programmable base editing of zebrafish genome using a modified CRISPR-Cas9 system. Nat. Commun. 8, 118 (2017).

Chadwick, A. C., Wang, X. & Musunuru, K. In vivo base editing of PCSK9 (proprotein convertase subtilisin/kexin type 9) as a therapeutic alternative to genome editing. Arterioscler. Thromb. Vasc. Biol. 37, 1741–1747 (2017).

Liu, Z. et al. Efficient generation of mouse models of human diseases via ABE- and BE-mediated base editing. Nat. Commun. 9, 2338 (2018).

Wu, Y. et al. Increasing cytosine base editing scope and efficiency with engineered Cas9-PmCDA1 fusions and the modified sgRNA in rice. Front. Genet. 10, 379 (2019).

Ren, B. et al. A CRISPR/Cas9 toolkit for efficient targeted base editing to induce genetic variations in rice. Sci. China Life Sci. 60, 516–519 (2017).

Lee, H. K. et al. Simultaneous targeting of linked loci in mouse embryos using base editing. Sci. Rep. 9, 1662 (2019).

Gapinske, M. et al. CRISPR-SKIP: programmable gene splicing with single base editors. Genome Biol. 19, 107 (2018).

Gehrke, J. M. et al. An APOBEC3A-Cas9 base editor with minimized bystander and off-target activities. Nat. Biotechnol. 36, 977–982 (2018).

Zhou, C. et al. Highly efficient base editing in human tripronuclear zygotes. Protein Cell 8, 772–775 (2017).

Yuan, J. et al. Genetic modulation of RNA splicing with a CRISPR-guided cytidine deaminase. Mol. Cell 72, 380–394.e7 (2018).

Hua, K., Tao, X. & Zhu, J.-K. Expanding the base editing scope in rice by using Cas9 variants. Plant Biotechnol. J. 17, 499–504 (2019).

Li, X. et al. Base editing with a Cpf1–cytidine deaminase fusion. Nat. Biotechnol. 36, 324–327 (2018).

Lee, C. et al. CRISPR-pass: gene rescue of nonsense mutations using adenine base editors. Mol. Ther. 27, 1364–1371 (2019).

Liu, Z. et al. Efficient base editing with expanded targeting scope using an engineered Spy-mac Cas9 variant. Cell Discov. 5, 58 (2019).

Ren, B. et al. Cas9-NG greatly expands the targeting scope of the genome-editing toolkit by recognizing NG and other atypical PAMs in rice. Mol. Plant 12, 1015–1026 (2019).

Zhong, Z. et al. Improving plant genome editing with high-fidelity xCas9 and non-canonical PAM-targeting Cas9-NG. Mol. Plant 12, 1027–1036 (2019).

Liu, Z. et al. Precise base editing with CC context-specificity using engineered human APOBEC3G-nCas9 fusions. BMC Biol. 18, 111 (2019).

Liu, Z. et al. Improved base editor for efficient editing in GC contexts in rabbits with an optimized AID-Cas9 fusion. FASEB J. 33, 9210–9219 (2019).

Endo, M. et al. Genome editing in plants by engineered CRISPR–Cas9 recognizing NG PAM. Nat. Plants 5, 14–17 (2019).

Richter, M. F. et al. Phage-assisted evolution of an adenine base editor with improved Cas domain compatibility and activity. Nat. Biotechnol. 38, 883–891 (2020).

Hu, Z. et al. A compact Cas9 ortholog from Staphylococcus Auricularis (SauriCas9) expands the DNA targeting scope. PLOS Biol. 18, e3000686 (2020).

Agudelo, D. et al. Versatile and robust genome editing with Streptococcus thermophilus CRISPR1-Cas9. Genome Res. 30, 107–117 (2020).

Tan, J., Zhang, F., Karcher, D. & Bock, R. Expanding the genome-targeting scope and the site selectivity of high-precision base editors. Nat. Commun. 11, 629 (2020).

Li, X. et al. Programmable base editing of mutated TERT promoter inhibits brain tumour growth. Nat. Cell Biol. 22, 282–288 (2020).

Wang, X. et al. Cas12a base editors induce efficient and specific editing with low DNA damage response. Cell Rep. 31, 107723 (2020).

Doman, J. L., Raguram, A., Newby, G. A. & Liu, D. R. Evaluation and minimization of Cas9-independent off-target DNA editing by cytosine base editors. Nat. Biotechnol. 38, 620–628 (2020).

Slaymaker, I. M. et al. Rationally engineered Cas9 nucleases with improved specificity. Science 351, 84–88 (2016).

Rees, H. A. et al. Improving the DNA specificity and applicability of base editing through protein engineering and protein delivery. Nat. Commun. 8, 15790 (2017).

Xu, W. et al. Multiplex nucleotide editing by high-fidelity Cas9 variants with improved efficiency in rice. BMC Plant Biol. 19, 511 (2019).

Lee, J. K. et al. Directed evolution of CRISPR-Cas9 to increase its specificity. Nat. Commun. 9, 3048 (2018).

Liang, P. et al. Effective gene editing by high-fidelity base editor 2 in mouse zygotes. Protein Cell 8, 601–611 (2017).

Kim, D., Kim, D.-E., Lee, G., Cho, S.-I. & Kim, J.-S. Genome-wide target specificity of CRISPR RNA-guided adenine base editors. Nat. Biotechnol. 37, 430–435 (2019).

Hong, R., Ma, S. & Wang, F. Improving the specificity of adenine base editor using high-fidelity Cas9. Preprint at bioRxiv (2019).

Casini, A. et al. A highly specific SpCas9 variant is identified by in vivo screening in yeast. Nat. Biotechnol. 36, 265–271 (2018).

Thuronyi, B. W. et al. Continuous evolution of base editors with expanded target compatibility and improved activity. Nat. Biotechnol. 37, 1070–1079 (2019).

Yu, Y. et al. Cytosine base editors with minimized unguided DNA and RNA off-target events and high on-target activity. Nat. Commun. 11, 2052 (2020).

Wang, X. et al. Efficient base editing in methylated regions with a human APOBEC3A-Cas9 fusion. Nat. Biotechnol. 36, 946–949 (2018).

Gaudelli, N. M. et al. Directed evolution of adenine base editors with increased activity and therapeutic application. Nat. Biotechnol. 38, 892–900 (2020).

Zhou, C. et al. Off-target RNA mutation induced by DNA base editing and its elimination by mutagenesis. Nature 571, 275–278 (2019).

Rees, H. A., Wilson, C., Doman, J. L. & Liu, D. R. Analysis and minimization of cellular RNA editing by DNA adenine base editors. Sci. Adv. 5, eaax5717 (2019).

Liu, Z. et al. Efficient base editing with high precision in rabbits using YFE-BE4max. Cell Death Dis. 11, 36 (2020).

Martin, A. S. et al. A panel of eGFP reporters for single base editing by APOBEC-Cas9 editosome complexes. Sci. Rep. 9, 497 (2019).

St. Martin, A. et al. A fluorescent reporter for quantification and enrichment of DNA editing by APOBEC–Cas9 or cleavage by Cas9 in living cells. Nucleic Acids Res. 46, e84 (2018).

Coelho, M. A. et al. BE-FLARE: a fluorescent reporter of base editing activity reveals editing characteristics of APOBEC3A and APOBEC3B. BMC Biol. 16, 150 (2018).

Zuo, E. et al. A rationally engineered cytosine base editor retains high on-target activity while reducing both DNA and RNA off-target effects. Nat. Methods 17, 600–604 (2020).

Zuo, E. et al. Cytosine base editor generates substantial off-target single-nucleotide variants in mouse embryos. Science 364, 289–292 (2019).

Grünewald, J. et al. CRISPR DNA base editors with reduced RNA off-target and self-editing activities. Nat. Biotechnol. 37, 1041–1048 (2019).

Wang, T., Badran, A. H., Huang, T. P. & Liu, D. R. Continuous directed evolution of proteins with improved soluble expression. Nat. Chem. Biol. 14, 972–980 (2018).

Rees, H. A. & Liu, D. R. Base editing: precision chemistry on the genome and transcriptome of living cells. Nat. Rev. Genet. 19, 770–788 (2018).

Shimatani, Z. et al. Targeted base editing in rice and tomato using a CRISPR-Cas9 cytidine deaminase fusion. Nat. Biotechnol. 35, 441–443 (2017).

Sasaguri, H. et al. Introduction of pathogenic mutations into the mouse Psen1 gene by Base Editor and Target-AID. Nat. Commun. 9, 2892 (2018).

Farzadfard, F. et al. Single-nucleotide-resolution computing and memory in living cells. Mol. Cell 75, 769–780.e4 (2019).

Shevidi, S., Uchida, A., Schudrowitz, N., Wessel, G. M. & Yajima, M. Single nucleotide editing without DNA cleavage using CRISPR/Cas9-deaminase in the sea urchin embryo. Dev. Dyn. 246, 1036–1046 (2017).

Banno, S., Nishida, K., Arazoe, T., Mitsunobu, H. & Kondo, A. Deaminase-mediated multiplex genome editing in Escherichia coli. Nat. Microbiol. 3, 423–429 (2018).

Wang, Y. et al. MACBETH: multiplex automated Corynebacterium glutamicum base editing method. Metab. Eng. 47, 200–210 (2018).

Xie, J. et al. Efficient base editing for multiple genes and loci in pigs using base editors. Nat. Commun. 10, 2852 (2019).

Zong, Y. et al. Efficient C-to-T base editing in plants using a fusion of nCas9 and human APOBEC3A. Nat. Biotechnol. 36, 950–953 (2018).

Gammage, P. A., Moraes, C. T. & Minczuk, M. Mitochondrial genome engineering: the revolution may not be CRISPR-ized. Trends Genet. 34, 101–110 (2018).

Molla, K. A. & Yang, Y. CRISPR/Cas-mediated base editing: technical considerations and practical applications. Trends Biotechnol. 37, 1121–1142 (2019).

Hess, G. T., Tycko, J., Yao, D. & Bassik, M. C. Methods and applications of CRISPR-mediated base editing in eukaryotic genomes. Mol. Cell 68, 26–43 (2017).

Liu, Z. et al. Highly efficient RNA-guided base editing in rabbit. Nat. Commun. 9, 2717 (2018).

Li, Q. et al. CRISPR-Cas9-mediated base-editing screening in mice identifies DND1 amino acids that are critical for primordial germ cell development. Nat. Cell Biol. 20, 1315–1325 (2018).

Yeh, W.-H., Chiang, H., Rees, H. A., Edge, A. S. B. & Liu, D. R. In vivo base editing of post-mitotic sensory cells. Nat. Commun. 9, 2184 (2018).

Zeng, Y. et al. Correction of the Marfan syndrome pathogenic FBN1 mutation by base editing in human cells and heterozygous embryos. Mol. Ther. 26, 2631–2637 (2018).

Li, G. et al. Highly efficient and precise base editing in discarded human tripronuclear embryos. Protein Cell 8, 776–779 (2017).

Liang, P. et al. Correction of β-thalassemia mutant by base editor in human embryos. Protein Cell 8, 811–822 (2017).

Zong, Y. et al. Precise base editing in rice, wheat and maize with a Cas9-cytidine deaminase fusion. Nat. Biotechnol. 35, 438–440 (2017).

Lu, Y. & Zhu, J. K. Precise editing of a target base in the rice genome using a modified CRISPR/Cas9 system. Mol. Plant 10, 523–525 (2017).

Park, D.-S. et al. Targeted base editing via RNA-guided cytidine deaminases in Xenopus laevis embryos. Mol. Cells 40, 823–827 (2017).

Rossidis, A. C. et al. In utero CRISPR-mediated therapeutic editing of metabolic genes. Nat. Med. 24, 1513–1518 (2018).

Kim, K. et al. Highly efficient RNA-guided base editing in mouse embryos. Nat. Biotechnol. 35, 435–437 (2017).

Tanaka, S. et al. In vivo targeted single-nucleotide editing in zebrafish. Sci. Rep. 8, 1–11 (2018).

Qin, L. et al. High-efficient and precise base editing of C•G to T•A in the allotetraploid cotton (Gossypium hirsutum) genome using a modified CRISPR/Cas9 system. Plant Biotechnol. J. 18, 45–56 (2020).

Chen, Y. et al. CRISPR/Cas9-mediated base-editing system efficiently generates gain-of-function mutations in Arabidopsis. Sci. China Life Sci. 60, 520–523 (2017).

Li, J., Sun, Y., Du, J., Zhao, Y. & Xia, L. Generation of targeted point mutations in rice by a modified CRISPR/Cas9 system. Mol. Plant 10, 526–529 (2017).

Li, Y. et al. Programmable single and multiplex base-editing in Bombyx mori using RNA-guided cytidine deaminases. G3 (Bethesda) 8, 1701–1709 (2018).

Qin, W. et al. Precise A•T to G•C base editing in the zebrafish genome. BMC Biol. 16, 1–8 (2018).

Li, C. et al. Expanded base editing in rice and wheat using a Cas9-adenosine deaminase fusion. Genome Biol. 19, 59 (2018).

Liang, P. et al. Effective and precise adenine base editing in mouse zygotes. Protein Cell 9, 808–813 (2018).

Ma, Y. et al. Highly efficient and precise base editing by engineered dCas9-guide tRNA adenosine deaminase in rats. Cell Discov. 4, 1–4 (2018).

Kang, B.-C. et al. Precision genome engineering through adenine base editing in plants. Nat. Plants 4, 427–431 (2018).

Hua, K., Tao, X., Yuan, F., Wang, D. & Zhu, J.-K. Precise A· T to G· C base editing in the rice genome. Mol. Plant 11, 627–630 (2018).

Song, C.-Q. et al. Adenine base editing in an adult mouse model of tyrosinaemia. Nat. Biomed. Eng. 4, 125–130 (2020).

Yan, F. et al. Highly efficient A· T to G· C base editing by Cas9n-guided tRNA adenosine deaminase in rice. Mol. Plant 11, 631–634 (2018).

Kuscu, C. et al. CRISPR-STOP: gene silencing through base-editing-induced nonsense mutations. Nat. Methods 14, 710–712 (2017).

Billon, P. et al. CRISPR-mediated base editing enables efficient disruption of eukaryotic genes through induction of STOP codons. Mol. Cell 67, 1068–1079.e4 (2017).

Tang, W. & Liu, D. R. Rewritable multi-event analog recording in bacterial and mammalian cells. Science 360, eaap8992 (2018).

Hwang, B. et al. Lineage tracing using a Cas9-deaminase barcoding system targeting endogenous L1 elements. Nat. Commun. 10, 1234 (2019).

Yin, K., Gao, C. & Qiu, J.-L. Progress and prospects in plant genome editing. Nat. Plants 3, 17107 (2017).

Anzalone, A. V. et al. Search-and-replace genome editing without double-strand breaks or donor DNA. Nature 576, 149–157 (2019).

Heyer, W.-D., Ehmsen, K. T. & Liu, J. Regulation of homologous recombination in eukaryotes. Annu. Rev. Genet. 44, 113–139 (2010).

Moynahan, M. E. & Jasin, M. Mitotic homologous recombination maintains genomic stability and suppresses tumorigenesis. Nat. Rev. Mol. Cell Biol. 11, 196–207 (2010).

Jiang, T. et al. Chemical modifications of adenine base editor mRNA and guide RNA expand its application scope. Nat. Commun. 11, 1979 (2020).

Zeng, J. et al. Therapeutic base editing of human hematopoietic stem cells. Nat. Med. 26, 535–541 (2020).

Ran, F. A. et al. Genome engineering using the CRISPR-Cas9 system. Nat. Protoc. 8, 2281–2308 (2013).

Chen, J. S. et al. Enhanced proofreading governs CRISPR–Cas9 targeting accuracy. Nature 550, 407–410 (2017).

Kim, J. et al. Structural and kinetic characterization of Escherichia coli TadA, the wobble-specific tRNA deaminase. Biochemistry 45, 6407–6416 (2006).

Gao, X. et al. Treatment of autosomal dominant hearing loss by in vivo delivery of genome editing agents. Nature 553, 217–221 (2018).

Ma, X. et al. Analysis of error profiles in deep next-generation sequencing data. Genome Biol. 20, 50 (2019).

Petrackova, A. et al. Standardization of sequencing coverage depth in NGS: recommendation for detection of clonal and subclonal mutations in cancer diagnostics. Front. Oncol. 9, 851 (2019).

Hwang, G.-H. et al. Web-based design and analysis tools for CRISPR base editing. BMC Bioinformatics 19, 542 (2018).

Clement, K. et al. CRISPResso2 provides accurate and rapid genome editing sequence analysis. Nat. Biotechnol. 37, 224–226 (2019).

Kluesner, M. G. et al. EditR: a method to quantify base editing from Sanger sequencing. CRISPR J. 1, 239–250 (2018).

Tsai, S. Q. et al. GUIDE-seq enables genome-wide profiling of off-target cleavage by CRISPR-Cas nucleases. Nat. Biotechnol. 33, 187–197 (2015).

Tsai, S. Q. et al. CIRCLE-seq: a highly sensitive in vitro screen for genome-wide CRISPR–Cas9 nuclease off-targets. Nat. Methods 14, 607–614 (2017).

Kim, D. et al. Digenome-seq: genome-wide profiling of CRISPR-Cas9 off-target effects in human cells. Nat. Methods 12, 237–243 (2015).

Gao, Z., Harwig, A., Berkhout, B. & Herrera-Carrillo, E. Mutation of nucleotides around the +1 position of type 3 polymerase III promoters: the effect on transcriptional activity and start site usage. Transcription 8, 275–287 (2017).

Kim, S., Bae, T., Hwang, J. & Kim, J.-S. Rescue of high-specificity Cas9 variants using sgRNAs with matched 5′ nucleotides. Genome Biol. 18, 218 (2017).

Labun, K., Montague, T. G., Gagnon, J. A., Thyme, S. B. & Valen, E. CHOPCHOP v2: a web tool for the next generation of CRISPR genome engineering. Nucleic Acids Res. 44, W272–W276 (2016).

Haeussler, M. et al. Evaluation of off-target and on-target scoring algorithms and integration into the guide RNA selection tool CRISPOR. Genome Biol. 17, 148 (2016).

Bae, S., Park, J. & Kim, J. S. Cas-OFFinder: a fast and versatile algorithm that searches for potential off-target sites of Cas9 RNA-guided endonucleases. Bioinformatics 30, 1473–1475 (2014).

Supplementary Material

The authors are grateful for the patients who provided blood for the previously mentioned studies, research support from the Leukemia & Lymphoma Society and the National Cancer Institute (P50 CA140158 and 1K12 <"type":"entrez-nucleotide","attrs":<"text":"CA133250","term_id":"35019051","term_text":"CA133250">> CA133250). The authors also thank Els Vanstreels for help with confocal imaging ( Figure 2 G-H).

This work is funded by the National Institutes of Health (R01-GM069909 Y.M.C.), Welch Foundation (I-1532 Y.M.C.), Leukemia & Lymphoma Society Scholar award (Y.M.C.), Cancer Prevention Research Institute of Texas (CPRIT RP120352 Y.M.C. and PR-101496 Q.S.), and University of Texas Southwestern Endowed Scholars Program (Y.M.C.). Results shown in this report are derived from work performed at Argonne National Laboratory, Structural Biology Center at the Advanced Photon Source. Argonne is operated by UChicago Argonne LLC, for the US Department of Energy, Office of Biologic and Environmental Research under contract DE-AC02-06CH11357. All the immunofluorescence images presented in this paper (except Figure 2 G-H) were collected at the Confocal Microscopy Imaging Facility (CMIF) at The Ohio State University. Mr and Mrs Michael Thomas, The Harry Mangurian Foundation, and The D. Warren Brown Foundation also supported this work.

Watch the video: Streak Plate Technique for The Isolation of Pure CultureA Complete Procedure Microbiology (May 2022).


  1. Barta

    What words ... Super

  2. Hwitcomb

    No, front.

  3. Einhard

    the quality is normal, I thought it would be worse, but I was wrong and I'm glad about it)

  4. Rez

    Great message, I like it :)

  5. Manus

    In my opinion, the topic is very interesting. I suggest you discuss it here or in PM.

Write a message