Construction and quantitation of a selectable protein splicing sensor using Gibson assembly and spot titers

Abstract

Inteins (intervening proteins) are translated within host proteins and removed through protein splicing. Conditional protein splicing (CPS), where the rate and accuracy of splicing is highly dependent on environmental cues, has emerged as a novel form of post-translational regulation. While CPS has been demonstrated for several inteins in vitro, a comprehensive understanding of inteins requires tools to quantitatively monitor intein activity within the cellular context. Here, we describe a method for construction of a splicing-dependent system that can be used to quantitatively assay for conditions that modulate protein splicing.

INTRODUCTION

Abundant and widespread in the microbial world, inteins (intervening proteins) have garnered interest as tools for biotechnological application (Wood et al., 2014; Sarmiento and Camarero, 2019), as agents of genome evolution (Gogarten and Hilario, 2006), and as antimicrobial targets (Chan et al. 2016). Inteins are mobile elements translated within host proteins and removed through protein splicing. In this process, the intein autocatalytically removes itself by breaking two peptide bonds and rejoining the flanking sequences (called N-and C-exteins) with a peptide bond.

This seamless ability to shuffle peptide bonds has proven exceptionally useful in protein engineering applications. Inteins can be thought of as self-contained, single-turnover enzymes with a unique ability to break and remake peptide bonds at designated positions. As such, this ability has been exploited in numerous protein engineering applications including bioseparations, bioconjugation, biosensing, and protein cyclization (Wood et al., 2014; Sarmiento and Camarero, 2019). The impact of intein-based technologies on protein engineering has been, and continues to be, profound.

The canonical mechanism of protein splicing, class 1, proceeds in four steps (Mills et al., 2014). In step 1, the first residue of the intein (known as the 1 position; either a Cysteine or Serine) performs a nucleophilic attack on the preceding peptide bond, forming a (thio)ester linkage. In step 2, the first residue of the C-extein (known as the +1 position; either a Cysteine, Serine, or Threonine) performs a nucleophilic attack on the (thio)ester resulting from step 1, forming a branched intermediate. In step 3, a conserved asparagine found at the end of the intein, cyclizes to liberate the intein.

Finally, in step 4, the (thio)ester linking the N- and C-exteins rearranges to form a peptide bond. Off pathway reactions can also occur, resulting in N- or C-extein cleavage from the intein prior to ligation. For example, following formation of this (thio)ester, the +1 nucleophile can attack, or the N-extein can be cleaved from the Intein-C-extein prior to ligation. Two alternative splicing mechanisms exist as well. Class 2 and 3 inteins use a similar strategy as class 1 inteins. For Class 2 inteins, there is no initiating nucleophile. For Class 3 inteins, the position of the initiating nucleophile is located internally rather than at the start of the intein sequence (Mills et al., 2014).

Inteins are present in the genomes of archaea, bacteria, unicellular eukaryotes, phages, and viruses (Novikova et al. 2016). Inteins are not only widespread and abundant in the microbial world, they come in different formats. Inteins most commonly house as homing endonuclease domains (HEN) between the splicing blocks. These HENs assist in the invasion of intein-free alleles, making some inteins mobile genetic elements. Mini-inteins, which lack HEN domains, contain little more than conserved sequences, known as splicing blocks, necessary for activity. Most inteins found in nature are contiguous, whether HEN containing or not, with the intein and the interrupted protein translated as a single polypeptide that undergoes cis splicing. However, a subset of inteins in nature have been split within the host genome, with the intein-containing gene expressed and translated independently as two halves that splice in trans upon reassembly.

Although traditionally considered parasitic elements (Naor et al., 2016), compelling examples of conditional protein splicing (CPS) in response to environmental stress have been recently discovered (Lennon et al., 2017). Understanding this emerging form of post-translational regulation requires tools to study CPS within living cells. Herein, we describe a strategy for construction of a splicing-dependent selectable marker that can be used to monitor intein activity within the cellular environment (Figure 1). This marker can be used to study stress conditions that block splicing, as well as any factors that may increase intein activity. As the presence of the intein likely disrupts host protein activity prior to splicing, regulation of protein splicing ultimately controls host protein function. We focus on a Mycobacterium smegmatis DnaB helicase intein 1 (Msm DnaBi1) within a kanamycin resistance marker (KanR2). M. smegmatis DnaB contains two inteins, designated DnaBi1 and DnaBi2. DnaBi1 is a 139-residue class 3 intein lacking a HEN domain, while DnaBi2 is 425-residue class 1 intein containing a HEN domain. Msm DnaBi1 was previously shown to be reversibly inhibited by both oxidative stress and zinc within M. smegmatis (Kelley et al., 2018; Woods et al., 2020). In principle, a splicing-dependent sensor can be developed with any combination of intein and selectable marker using the techniques described within this protocol. The first protocol details methodology to construct a series of DnaBi1-KanR2 fusions using Gibson Assembly. The purpose of this protocol is to provide a library of intein-marker fusions to screen in the second protocol. This second protocol describes the process of screening the DnaBi1-KanR2 fusion library using quantitative spot titers to determine which with intein-marker fusions provide splicing-dependent resistance. The purpose of protocol is to find an intein-marker fusion that requires splicing to provide resistance, as well as to quantitatively measure protein splicing levels directly in the cellular environment. We also provide a support protocol for preparation of competent cells and transformation using M. smegmatis for studies of mycobacterial inteins.

Figure 1:

Design of KanR2-Msm DnaB1 intein fusion. Splicing-dependent resistance phenotype depicted. Splicing-independent resistance and no resistance phenotypes are also possible and described in Basic Protocol 2 and Commentary sections. N-extein, blue; Intein, red; Intein initiating nucleophile C118, yellow; C-extein, green; +1 serine of of C-extein, purple.

BASIC PROTOCOL 1

Construction of an Intein-containing KanR2 Library using Gibson Assembly

This protocol is used to build a series of constructs with an intein inserted throughout the primary sequence of a genetic marker. For this example, Msm DnaBi1, both wildtype and a catalytically inactive version of the intein (C118), is cloned immediately upstream of all serine residues (the natural +1 nucleophile) within KanR2 using Gibson assembly. For splicing to precede, the only strict requirement is a +1 nucleophile (serine, cysteine, or threonine) as the first residue of the C-extein. The purpose of this protocol is to construct a library that can be screened for splicing-dependent resistance SDR) in Basic Protocol 2.

Materials

pUC4K plasmid (GE Healthcare or available from authors upon request)

Msm DnaBi1-containing plasmid (available from authors upon request)

2X CloneAmp PCR Premix (Takara #RR030A)

Gibson Assembly Master Mix (New England Biolabs #E2611)

NEB5α chemically competent Escherichia coli cells (New England Biolabs # C2987H)

LB agar (Research Products International # L24030)

Carbenicillin (Gold Biotechnology #C-103)

Gene specific primers from commercial vendors (Integrated DNA Technologies)

DNA clean and concentrator kit (Zymo Research # D4004)

DNA miniprep kit (Omega Bio-Tek # D6942-01)

PCR tubes (Thermo Scientific AB-1182)

Roll and Grow plating beads (MP Bio #115000550-CF)

Petri plates (Fisher # 08-757-105)

Nanodrop Spectrophotometer (Thermo Fisher #ND-2000 or equivalent)

Thermocycler

Ice

Ice Bucket

1. Design primers to amplify intein and vector backbone for Gibson assembly using PCR.

   1. NEB Builder 2.0 (https://nebuilder.neb.com/#!/) can be used to design primers.
   2. An example of primers used for amplification is shown in Figure 2. We abbreviate the fusions as KD (KanR2-DnaBi1) followed by the position of the Ser +1 residue and whether it is the catalytically active (WT) or inacative (MUT) intein. For example, for the catalytically active intein inserted between residues 153 and 154 of KanR2 where Ser154 is utilized as the +1 residue, the construct is referred to as KD154:WT. The cloning scheme for KD154:WT is shown in Figure 2.

2. Synthesize primers using a commercial company such as Integrated DNA technologies (IDT).

3. Amplify PCR products for Gibson assembly using 2X CloneAmp PCR Premix

   1. Thaw all components on ice and mix the following:

      1. 12.5 μL 2X CloneAmp PCR Premix
      2. 0.25 pmol of each primer
      3. 50 ng template DNA
      4. ddH₂O to a final volume of 25 μL
   2. Use the following amplification conditions in a thermocycler. Allow for 30 total cycles

      1. 98°C for 10 seconds
      2. 55°C for 15 seconds
      3. 72°C for 5 seconds per kb
   3. The desired PCR products include:

      1. Vector backbone using inverse PCR
         Inverse PCR will serve to linearize the vector backbone without the use of restriction endonucleases.
      2. WT Intein (Int_WT) using PCR
      3. Catalytically inactive intein (Int_MUT) using PCR
         Msm DnaBi1 is a class 3 intein with an internal initiating nucleophile. Because of this, forward and reverse primers will be the same for both the WT and catalytically inactive intein (C118A). For class 1 inteins, a different forward primer is required as the initiating nucleophile is the first residue of the intein.

4. Purify PCR products using a DNA clean and concentrator kit.

5. Measure the concentration of the DNA using a spectrophotometer.

6. Perform Gibson Assembly reaction according to New England Biolabs specifications. For calculation of ng DNA needed, use NEBioCalculator dsDNA:

   Mass to/from Moles Convertor (http://nebiocalculator.neb.com/#!/dsdnaamt).

   1. Thaw all components on ice and mix the following:

      1. 10 μL 2X Gibson Assembly Master Mix
      2. 1–2 μL (0.1 pmol) linearized vector backbone

         1. For pUC4K, 242 ng
      3. 1–2 μL (0.2 pmol) Int_WT DNA or Int_MUT DNA

         1. For dnaBi1, 52 ng
      4. ddH₂O to a final volume of 20 μL
   2. Incubate at 50°C for 15 minutes

7. For each reaction, thaw 50 μL of chemically competent NEB 5α E. coli cells on ice for 10 minutes.

8. Gently mix 2 μL of the reaction from step 7 with 50 μL of chemically competent NEB 5α E. coli cells

9. Perform transformation of NEB 5α E. coli cells.

   1. Incubate on ice for 30 minutes
   2. Heat shock at 42°C for 30 seconds
   3. Incubate on ice for 5 minutes
   4. Add 950 μL of LB broth
   5. Shake at 250 RPM at 37°C for 1 hour. During this incubation period, prewarm three LB agar plates containing 50 μg/mL carbenicillin for each transformation.
   6. Place ~10–15 sterile plating beads on one LB agar plate with 50 μg/mL carbenicillin

      1. Select for resistance to carbenicillin, not kanamycin
   7. Add 200 μL of the transformation mix
   8. Spread cells over plate using plating beads
   9. Transfer plating beads to second plate and spread cells

      • i.
        Invert first plate directly onto second plate and gently tap to transfer beads
      • j.
        Transfer plating beads to third plate and spread cells. Dispose of beads.

10. Incubate at 37°C between 16–20 hours

11. With a single, isolated colony, inoculate 5 mL of LB media containing 50 μg/mL carbenicillin.

    1. For each separate transformant, it is suggested that three colonies be selected and prepared for downstream sequencing.

12. Grow these cultures to saturation shaking at 37° C overnight between 12–16 hours.

13. Purify plasmids using DNA miniprep kit.

14. Verify correct clones by DNA sequencing. Primers from the Gibson assembly reaction may be used for this purpose.

Figure 2:

KanR2-Msm DnaB1 intein fusion cloning. (A) Overall Gibson assembly cloning strategy for seamless WT Msm DnaB1 intein insertion between codons 153 and 154 of KanR2 (i.e. KD154:WT construction) example is shown. (B) Primers and Msm DnaBi1 and KanR2 DNA sequences for KD154:WT construction. For panel A, N-extein, blue; Intein, red; Intein initiating nucleophile C118, yellow; C-extein, green; For panel B. colored as in panel A, with intein initiating nucleophile C118 colored yellow and +1 serine of C-extein colored purple. Underlined regions indicate where primers anneal during PCR amplification.

BASIC PROTOCOL 2

Quantitative Spot Titers and Phenotype Determination

This protocol describes how to grow, dilute, and spot E.coli for quantitative determination of colony forming units (CFUs) over a range of kanamycin concentrations. The purpose of the quantitative spot titers is to determine the level of kanamycin resistance for each KanR2-intein fusion and if intein splicing from KanR2 is required for resistance. A similar approach can be useful to examine splicing within the native mycobacterial host of the intein, as well as be combined with different stress conditions (see Understanding Results; Kelley et al., 2018; Woods et al., 2020).

Materials

pUC18 plasmid (Addgene # 50004)

LB agar (See recipe)

LB agar plates with carbenicillin (See recipe)

LB agar plates with carbenicillin and variable kanamycin (See recipe)

LB medium (See recipe)

Disposable pipette tips (USA Scientific, Inc. Tip One or equivalent)

50 mL conical tubes (Corning # 352070)

Disposable cuvettes (Bio-Rad #2239950)

Spectrophotometer (Bio-Rad SmartSpec Plus Spectrophotometer or equivalent)

Multichannel pipettes (Thermo Fisher Scientific #46300000, 46300200, 46300400 or equivalent)

Microwave

Water bath at 55°C

Quantitative Spot Titers

1. Inoculate 5 mL of LB media containing the 50 μg/mL carbenicillin with a single colony of E.coli containing pUC4K (uninterrupted KanR2), pUC18 (kanamycin senstivite), or the intein-KanR2 fusions (INT_WT and INT_MUT for each position).

2. Grow these cultures to saturation shaking at 37°C overnight between 12–16 hours.

3. Determine the optical density at 600 nm (OD₆₀₀) of the cultures the following day by diluting 0.1 mL of the overnight (O.N.) culture into 0.9 mL of fresh LB medium.

4. Mix well and pipette into spectrophotometric cuvette.

5. Record the OD₆₀₀ and dilution ratio to make a 1mL at 1.0 OD₆₀₀.

   1. For example, if the culture is at OD₆₀₀ = 5, mix 200 μL culture with 800 μL fresh LB
6. Pipette 200 μL of 1.0 OD₆₀₀ (1×10⁰ OD₆₀₀) of each strain being tested into the first row of wells in a 96-well plate. Cover plate when not pipetting.

   1. Figure 3 shows an example arrangement for Kan + (pUC4K) in well A1, Kan – (pUC18) in well A2, KD116:WT in well A3, KD116:MUT in well A4, KD154:WT in well A5, KD154:MUT in well A6, KD189:WT in well A7, KD189:MUT in well A8, KD200:WT in well A9, and KD200:MUT in well A10

7. Aseptically decant fresh LB into an empty rectangular plate until full. This media will serve as the source of media for all dilutions.

8. Use the P1000 multichannel pipette to dispense 180 μL of fresh media into five rows of the 96-well plate below each strain to be tested.

   • a
     For Figure 3 example, rows B1–10, C1–10, D1–10, E1–10, and F1–10

9. Pipette 20 μL of culture with the P200 multichannel pipette from the top row of wells containing 1×10⁰ OD₆₀₀ cultures (Figure 3 A1–10) and dispense into the next row of wells (Figure 3 B1–10).

10. Mix 10 times using the multichannel pipette to make 1×10⁻¹ OD₆₀₀ dilution and discard tips.

11. Pipette 20 μL of 1×10⁻¹ OD₆₀₀ cultures (Figure 3 B1–10) with the multichannel pipette and dispense into the subsequent row (Figure 3 C1–10).

12. Mix 10 times using the multichannel pipette to make 1×10⁻² OD₆₀₀ dilution and discard tips.

13. Pipette 20 μL of 1×10⁻² OD₆₀₀ cultures (Figure 3 C1–10). with the multichannel pipette and dispense into the subsequent row (Figure 3 D1–10).

14. Mix 10 times using the multichannel pipette to make 1×10⁻³ OD₆₀₀ dilution and discard tips.

15. Pipette 20 μL of 1×10⁻³ OD₆₀₀ cultures (Figure 3 D1–10). with the multichannel pipette and dispense into the subsequent row (Figure 3 E1–10).

16. Mix 10 times using the multichannel pipette to make 1×10⁻⁴ OD₆₀₀ dilution and discard tips.

17. Pipette 20 μL of 1×10⁻⁴ OD₆₀₀ cultures (Figure 3 E1–10). with the multichannel pipette and dispense into the subsequent row (Figure 3 F1–10).

18. Mix 10 times using the multichannel pipette to make 1×10⁻⁵ OD₆₀₀ dilution and discard tips.

    • b
      At this point, each strain will have titers of 1×10⁰, 1×10⁻¹, 1×10⁻², 1×10⁻³, 1×10⁻⁴, and 1×10⁻⁵ OD₆₀₀ positioned in the 96-well plate (Figure 3).

19. Rotate the 96-well plate so that the serial dilutions are decreasing in OD₆₀₀ value, with the 1×10⁰ OD₆₀₀ sample on the left and the 1×10⁻⁵ OD₆₀₀ sample on the right.

20. Lay out the six in increasing kanamycin concentrations on the bench.

21. Use a P10 multichannel pipette to dispense 1.25 μL of each dilution for the first strain onto each of the plates (Figure 3). Be careful to spot the titers onto the same position of each plate and not to pierce the agar with the pipette tips. Also note that two positions of the multichannel pipette will not contain liquid.

22. Repeat step 21 for each of the strains being tested. Each subsequent strain should be spotted below the last (Figure 3). Ensure that spots between strains do not touch.

23. Cover the plates loosely for 20 minutes to allow ventilation and drying of the spots.

24. Secure the plate tops to the plates, invert, stack, and incubate 12–16 hours at 37°C.

25. Move plates from the incubator to the bench the following morning.

26. Make sure the positive control (KanR) grows on all kanamycin concentrations and that the negative control (KanS) only grows on plate without kanamycin.

    Note: KanR and KanS should exhibit growth to approximately the same dilution in the absence of kanamycin (Figure 4A).

27. Record the number of colonies formed at the highest dilution factor for which individual colonies can be counted. Do this for each strain at all kanamycin concentrations tested.

28. To calculate CFUs/mL:

    1. (# colonies) × (dilution factor) × (mL to μL conversion)

       1. mL conversion to μL spotted = 1000 μL /1.25 μL = 800
    2. For example, if five colonies are formed at a dilution factor of 1×10⁻³

       1. Five colonies (5) × dilution factor (10³) × mL to μL conversion (800) = 4 × 10⁶ CFUs/mL

29. Data may be organized in a table and plotted on a semi-log plot with the concentration of kanamycin on the x-axis (linear) and CFUs/mL on the y-axis (log10). CFUs should be calculated for each strain at each concentration of kanamycin.

    An example of SDR (KD154) and SIR (KD189) phenotypes are plotted in Figure 4B. For ease in visualization as the value zero cannot be plotted on, represent no colonies as 0.1 CFUs.

30. Determine resistance phenotype to each strain based on SDR, splicing-independent resistance (SIR), or no resistance (NR). All strains should exhibit growth to approximately the same dilution in the absence of kanamycin (Figure 4A).

    1. SDR phenotype: only catalytically active (Int_WT) intein-KanR2 fusions will grow in the presence of kanamycin. Catalytically inactive (Int_MUT) inteinKanR2 fusions will not grow in the presence of any kanamycin concentration tested (Figure 4A and 4B).
    2. SIR phenotype: both catalytically active (Int_WT) and inactive (Int_MUT) inteinKanR2 fusions will grow to the same dilution in the presence of kanamycin (Figure 4A and 4B).
    3. NR phenotype: both catalytically active (Int_WT) and inactive (Int_MUT) inteinKanR2 fusions will not grow in the presence of any concentration of kanamycin (Figure 4A).

Figure 3:

Preparation of strain dilutions and spot titer strategy. (A) Preparation of strain dilutions as described in Basic Protocol 2. For each strain indicated above columns 110, serial dilutions from OD₆₀₀ 10⁰ - 10⁻⁵ are made in rows A - F. (B) Scheme for spotting strain dilutions. Approximate positions of where indicated wells should be spotted using a multichannel pipette. Dilution spotting should be repeated for all kanamycin-containing plates. Note that only 6 of the 8 pipette tips contain cells. Strain positions correspond to example given in Figure 4. Images were generated using Biorender

Figure 4:

Representative Escherichia coli spot titer assays and semi-log plot of kanamycin concentration vs log₁₀ CFUs/mL. (A) Each E. coli strain contains either uninterrupted KanR2 (Kan +), no KanR2 (Kan −), or a single KanR2-Msm DnaBi1 fusion as indicated. Strains were spotted at indicated dilution. SIR, splicing-independent resistance; SDR, splicing-dependent resistance; NR, no resistance. Kanamycin concentration is 100 μg/mL when present. Positions of strains on plates correspond to positions in Figure 3. (B) Semi-log plot of representative data set with the concentration of kanamycin on the x-axis and vs log₁₀ CFUs/mL on the y-axis. Examples of SDR (KD154:WT vs. KD154:C118A) and SIR (KD189:WT vs. KD189:C118A) phenotypes are shown.

SUPPORT PROTOCOL 1

Preparation of Competent Cells and Transformation in Mycobacterium smegmatis

This support protocol describes the preparation of competent cells and transformation using M. smegmatis (van Kessel and Hatfull, 2008; Singh and Reyrat, 2009), the species which houses Msm DnaBi1. To monitor splicing in M. smegmatis, the SDR fusion determined in Basic Protocol 2 should be cloned into a mycobacterial shuttle vector using the Gibson assembly strategy described in Basic Protocol 1. Splicing can be measured with and without stress (Kelley et al., 2018; Woods et al., 2020) and as discussed in the Commentary.

Materials

M. smegmatis mc² 155 (available from authors upon request)

Plasmid DNA (pMBC283 or other mycobacterial shuttle vector; available from authors upon request)

Middlebrook 7H9 medium (See recipe)

Middlebrook 7H10 plates (See recipe)

10% sterile glycerol (See recipe)

Roll and Grow plating beads (MP Bio #115000550-CF)

Carbenicillin (Gold Biotechnology #C-103)

1.5 mL Eppendorf tubes (USA Scientific #4036-3204 or equivalent)

Culture tubes (VWR 60819–820 or equivalent)

Bio-Rad Electroporation Cuvettes, 0.2 cm gap (Bio-Rad # 1652082)

Bio-Rad Gene Pulser (Bio-Rad #1652660)

Sterile 500 mL baffled Erlenmeyer flask (Millipore Sigma #CLS4450500 or equivalent)

Refrigerated centrifuge (Eppendorf 5810R or equivalent)

Paper towel

Ice

Part 1: Preparation of M. smegmatis Competent Cells

• 1Place 10% sterile glycerol at 4°C.
• 2Inoculate 5 mL sterile Middlebrook 7H9 liquid medium with M. smegmatis mc² 155.
• 3Grow cells at 37°C to saturation. This will take approximately 2 days.
• 4Subculture cells into 100 mL fresh 7H9 medium in a 500 mL baffled flask to an OD₆₀₀ of 0.02.

  This protocol describes working with a 100 mL culture that will yield approximately 40 tubes of 100 μL competent cells. However, any volume of cells can be used, but keep the ratio of final competent cells consistent.

• 5Incubate cells at 37°C overnight to a final OD₆₀₀ of 0.8 – 1.0. Proceed to electrocompetent cell preparation. Chill M. smegmatis cells by placing the flask on ice for 30 minutes.

  Cells can be incubated at this step on ice for up to 2 hours if convenient.

• 6Depending on equipment, cells can either be transferred to a single container that can accommodate 100 mL volume or cells can be split into two 50 mL conical tubes. The steps described here will present the method for a single container approach.
• 7Centrifuge cells at 4°C at 3,600 x g for 10 minutes. Make sure to balance centrifuge appropriately if using a single container.
• 8Discard the supernatant into an appropriate waste container, careful to not disturb the pellet.
• 9Use a 50 mL serological pipet to carefully wash and resuspend the cell pellet in 50 mL, or 1/2 the initial culture volume, 10% sterile, 4°C glycerol. An aggressive resuspension can lead to loss of cell viability and reduced competency.
• 10Centrifuge to pellet the cells as described in Step 7 and discard the supernatant as in Step 8.
• 11Carefully, wash and resuspend the cell pellet in 25 mL, or 1/4 the initial culture volume, 10% sterile, 4°C glycerol.
• 12Centrifuge to pellet the cells as described in Step 7 and discard the supernatant as in Step 8.
• 13Carefully, wash and resuspend the cell pellet in 12.5 mL, or 1/8 the initial culture volume, 10% sterile, 4°C glycerol.
• 14Centrifuge to pellet the cells as described in Step 7 and discard the supernatant as in Step 8.
• 15Carefully, wash and resuspend the cell pellet in 10 mL, or 1/10 the initial culture volume, 10% sterile, 4°C glycerol.
• 16Centrifuge to pellet the cells as described in Step 7 and discard the supernatant as in Step 8.
• 17Carefully, resuspend the cell pellet in 4 mL, or 1/25 the initial culture volume, 10% sterile, 4°C glycerol. Pipette 100 μL of cells into sterile 1.5 mL microcentrifuge tubes, keeping on ice. Repeat to make approximately 40 aliquots. For long term, store at −80°C. Cells can also be used immediately for transformation.

Part 2: Plasmid Transformation

• 18For transformation, create a M. smegmatis program on the Bio-Rad Gene Pulser with the following settings: 2.5 kV, 1000 Ω, 25 μF. This can be saved onto the Bio-Rad Gene Pulser for future use.
• 19If not using freshly-made electrocompetent cells, place cells on ice and thaw for 10 minutes. One tube of cells (100 μL) per transformation is needed.
• 20While cells are thawing, place an equal number of cuvettes on ice to chill.
• 21Pipette 1 μL plasmid DNA (~5–50 ng) into cells and gently pipette or flick to mix.
• 22Let cells with DNA incubate on ice for 10 minutes.
• 23Transfer the entire cell-DNA volume into a chilled cuvette. Gently, tap the cuvette on the benchtop to remove any bubbles, which can lead to arcing, and ensure the cells are settled in the bottom of the cuvette.
• 24Wipe the cuvette off with a paper towel to remove any moisture from chilling on ice and place in the electroporator holder.
• 25Electroporate the cells using the M. smegmatis program created in Step 1. If the cells arc, discard the cells and repeat the transformation, starting at Step 2.
• 26After successful electroporation, add 1 mL Middlebrook 7H9 medium prewarmed to 37°C to the cells and transfer into a sterile culture tube.
• 27Let the cells recover by incubating at 37°C at 250 RPM for 2 hours.
• 28During the 2-hour recovery period, remove an appropriate number of selective 7H10 plates from the fridge and allow to warm up to room temperature on the benchtop.
• 29Plate transformations on selective 7H10 agar plates and incubate at 37°C for three to five days.

REAGENTS AND SOLUTIONS

Kanamycin stock solution (50 mg/mL)

For 10 mL, dissolve 500 mg of kanamycin sulfate (Thermo Fisher Scientific #11815024) in ddH₂O to a total volume of 10 mL

Pass through 0.22 μm filter to sterilize

Aliquot and store up to 3 months at −20°C

Carbenicillin stock solution (50 mg/mL)

For 10 mL, dissolve 1 g of carbenicillin (Gold Biotechnology #C-103) in ddH₂O to a total volume of 10 mL

Pass through 0.22 μm filter to sterilize

Aliquot and store up to 3 months at −20°C

LB medium

For 1 L, dissolve 20 g LB (Research Products International #L24060) in ddH₂O to a total volume of 1 L

Autoclave to sterilize

Store up to 6 weeks at 4°C

LB agar

For 1 L, dissolve 35 g LB agar (Research Products International # L24030) in ddH₂O to a total volume of 1 L

Autoclave to sterilize

Store up to 6 weeks at 4°C

LB agar plates with carbenicillin

Microwave 1L of LB agar with the lid on, partially unscrewed, in 30–45 second pulses interrupted with vigorous swirling until the agar is completely melted and the solution is liquid.

Caution: The melting temperature for agar is 85° C and it will readily burn skin when liquified.

Microwaving for longer periods of time can cause the lid to burst off of the bottle with some of the media bubbling over risking contamination and altering the volume of nutrients and agar in the media. If this happens carefully clean the spilled media and start over.

Keep the agar melted by placing the bottle in a 55° C water bath and allow to come down to temperature.

Once at 55° C, add 1000 μL of 50 mg/mL carbenicillin stock solution. Gently mix to avoid bubbles and pour agar into Petri plates.

LB agar plates with carbenicillin and variable kanamycin

Melt LB agar as directed under LB agar plates with carbenicillin

Lay out six Rectangular, single-well dishes (Thermo Fisher Scientific #267060), oriented in the same direction, with their top down on the bench

Label the bottom of the plate in the top right corner with the final concentration of kanamycin in μg/mL that each respective plate will contain. Each plate will contain 0, 12.5, 50, 100, 200, or 500 μg/mL kanamycin.

Flip the plates so they are bottom down and in increasing order with respect to kanamycin concentration.

Place six 50 mL conical tubes in a rack and label them with respective kanamycin concentrations.

Pipette the appropriate amount of kanamycin stock solution (see recipe) into the five respective 50 mL conical tubes. Note that no kanamycin is added for 0 μg/mL.

1. For a final concentration in μg/mL, add that many μL of kanamycin stock solution. For example, for a final concentration of 100 μg/mL, add 100 μL of kanamycin stock solution.

2. For consistency of results, it is important that the kanamycin stock be fresh and not freeze/thawed more than once.

Remove the lids of the plates and place them to the side.

Remove the bottle of melted LB agar from the water bath and add 1000 μL carbenicillin stock solution (see recipe).

Screw the lid back on and mix completely by rolling and swirling, without creating bubbles.

Aseptically decant 50 mL of LB agar with 50 μg/mL carbenicillin into the conical tubes for 0 and 12.5 μg/mL kanamycin.

Secure cap on conical tubes and mix by inverting five times. Avoid air bubbles.

Pour the 50 mL of media into the appropriately labelled plate. If any air bubbles form, push to the sides of the rectangular plate using a sterile pipette tip.

Repeat steps previous three steps until all plates are poured. move quickly to avoid LB agar solidification prior to mixing with kanamycin.

Place the lids loosely on the plates without covering completely to allow for air drying and reducing condensation.

Allow agar to solidify and cool before securing the top of the plate and rest plates at room temperature top-down overnight.

10% sterile glycerol

For 100 mL, measure 90 mL of ddH₂O into a graduated cylinder

Add 10 mL of 100% glycerol (Fisher scientific #BP229–1)

Cover top of cylinder with parafilm or plastic wrap and invert several times to mix thoroughly

Pass through 0.22 μm filter to sterilize

Store up to 6 weeks at 20–22°C

50% sterile glycerol

For 100 mL, measure 50 mL of ddH₂O into a graduated cylinder

Add 50 mL of 100% glycerol (Fisher scientific #BP229–1)

Cover top of cylinder with parafilm or plastic wrap and invert several times to mix thoroughly

Pass through 0.22 μm filter to sterilize

Store up to 6 weeks at 20–22°C

20% Tween 80

For 50 mL, add 10 mL Tween 80 (Fisher Scientific #T164–500) to a 50 mL conical tube

Add 40 mL ddH₂O

Incubate at 55°C for 30 minutes, inverting several times every 5 minutes, to solubilize

Pass through 0.22 μm filter to sterilize

Middlebrook 7H9 medium

Dissolve 4.7g Middlebrook 7H9 (BD Difco #271310) in ddH₂O to a total volume of 900 mL

Add 100 mL ADC enrichment solution (BD Difco #212352)

Add 10 mL 50% sterile glycerol

Add 2.5 mL 20% Tween 80

Pass through 0.22 μm filter to sterilize

Store up to 6 weeks at 4°C

Middlebrook 7H10 agar plates

Dissolve 19 g Middlebrook 7H10 agar (BD Difco #262710) in ddH₂O to a total volume of 900 mL

Autoclave to sterilize

Allow to cool to 55°C in water bath

Aseptically add 100 mL OADC enrichment (BD Difco #212240)

Aseptically add 10 mL 50% sterile glycerol

Aseptically add appropriate antibiotic

Mix carefully so as to avoid bubbles

Pour plates (Fisher Scientific #09-720-500)

Store up to 6 weeks at 4°C

COMMENTARY

Background Information

When present and active, the HEN domain allows an intein to spread at the DNA level. As such, inteins have been traditionally viewed as selfish genetic elements, the invasiveness of which helps remodel genomes (Gogarten and Hilario, 2006; Naor et al., 2016). We and others have challenged this model of pure selfishness and argue that many inteins, particularly those that have lost their HEN domain, have transitioned to benefit the host. Several studies have provided compelling examples of CPS, whereby splicing is highly dependent on environmental conditions. These conditions, often crucial to the survival of the host organisms and/or relevant to function of the intein-containing protein, include pH, redox, reactive oxygen/nitrogen species (ROS/RNS), salt, temperature, and even DNA damage (Mills et al., 2001; Callahan et al., 2011; Topilina et al., 2015a; Topilina et al., 2015b; Reitter et al., 2016; Ciragan et al., 2016; Lennon et al., 2016; Lennon et al., 2018; Kelley et al., 2018; Lennon et al. 2019; Green et al., 2019; Woods et al., 2020). Further, inteins can be engineered to respond to a variety of stimuli to control ligation or cleavage reactions (Buskirk et al., 2004; Skretas et al, 2005; Lockless and Muir, 2009; Peck et al., 2011), suggesting nature could also employ the same tactic. This growing body of evidence that some inteins can speed up or slow down protein splicing in response to stress, ultimately controlling active levels of the intein-housing protein, compels us to rethink the role of these elements in nature. As more and more examples of conditional protein splicing from natural systems emerge, tools such as the DnaBi1-KanR2 sensor described in this protocol can be used to understand the factors that control protein splicing within the cellular environment.

Inteins are found within essential genes of several human pathogens and are absent in metazoans, making them attractive drug targets. Toward this end, the anticancer drug cisplatin has been shown to inhibit the growth of Mycobacterium tuberculosis in an intein-specific manner (Zhang et al., 2011), and was shown by crystallography to bind directly to catalytic residues of the RecA intein (Chan et al., 2016). Additionally, recent evidence suggests that cisplatin also targets an intein within the essential splicesosomal Prp8 protein of the pathogen Cryptococcus neoformans (Li et al., 2019). Further, zinc has been shown to reversibly inhibit splicing of the DnaB helicase intein of the pathogen Mycobacterium leprae (Woods et al. 2020). Therefore, understanding the conditions that control the splicing of inteins from pathogens within the native host environment, which our approach can be used to study, could not only yield new insights into their response to stress, but lead to intein-specific inhibitors that selectively kill them.

Critical Parameters

Basic protocol 1: In construction of the selectable maker-intein fusions, the PCR step prior to Gibson assembly is critical. To ensure success, careful primer design is needed. We suggest using NEB Builder 2.0 (https://nebuilder.neb.com/#!/).

Basic protocol 2: In obtaining a fusion with a splicing-dependent resistance (SDR) phenotype, we suggest screening numerous intein insertion sites. Previous work has shown that distance from the active site can be used as predictor of SDR (Lockless and Muir, 2009; Shen et al., 2012; Apgar et al., 2012; Davis et al., 2015; Woods et al., 2020), with insertion closer to the active site increasing the likelihood of SDR. However, different inteins have demonstrated different position-dependent phenotypes within the same sensor protein (Woods et al. 2020; Lockless and Muir 2009). Using an unbiased screen of all potential +1 nucleophiles also provides the advantage of isolating the most dramatic SDR phenotype (i.e. no resistance for splicing inactive mutant and highest resistance to kanamycin following splicing).

Support Protocol 1: It should be noted that cells should be kept cold during this protocol. This includes using ice-cold glycerol, a refrigerated centrifuge at 4°C, and keeping tubes on ice as much as possible.

Troubleshooting

Basic protocol 1: If Gibson assembly fails to produce correct clones, verify that PCR products are the expected size using agarose gel electrophoresis. Products will both be linear run as predicted size. We found that if correctly sized PCR products are obtained, Gibson assembly is efficient and accurate in all attempts.

Basic protocol 2: If a splicing-dependent resistance (SDR) phenotype is not isolated using the native +1 residue of the intein, consider sensor-intein fusions utilizing alternative +1 residues. For example, if the native +1 residue is a serine, consider cysteine or threonine as the +1.

Understanding results

The overall goal of this protocol is to build a sensor that directly links protein splicing to sensor activation. In this protocol, we describe insertion of the Class 3 DnaB1 intein from M. smegmatis throughout KanR2 (the host protein), an aminoglycoside Ophosphotransferase APH(3=)-Ia that provides resistance to the antibiotic kanamycin. It is important to note that not all sites of intein insertion require splicing for activation of the host protein (Lockless and Muir, 2009; Shen et al., 2012; Davis et al., 2015; Woods et al., 2020). It is therefore important to screen several sites, although proximity to the active site can increase the likelihood of splicing-dependent host protein activation (Apgar et al., 2012; Woods et al. 2020).

For our DnaBi1-KanR2 fusions, when challenged with increasing kanamycin concentrations, three phenotypes present based on insertion position. For strains demonstrating splicing-dependent resistance (SDR), the splicing-inactive (C118A for DnaBi1) fusion will be sensitive to the lowest kanamycin concentration (e.g. 12.5 μg/mL kanamycin) tested, while the WT fusion will be resistant to kanamycin (Figure 4; KD154). The degree of resistance for a given SDR fusion will be based on the position within KanR2 where the intein is inserted. For the splicing-independent resistance (SIR), both the WT and splicing-inactive fusions will display approximately equal resistance to kanamycin (Figure 4; KD116 and KD189). For the no resistance (NR) phenotype, to both WT and splicing-inactive fusions will lack any resistance to kanamycin (Figure 4; KD200)

SDR fusions can also be used in the native intein-housing organism, in this case M. smegmatis, given the activity of the host protein can be screened or selected using genetics. If utilizing intein-KanR2 fusions to examine conditional protein splicing, it is critical to use kanamycin and treatment conditions that do not lead to reduced survival alone, but are sufficiently high to ensure that reduction in KanR levels due to splicing inhibition would be detectable. If zinc, for example, were used to probe for in vivo splicing inhibition, significant differences in survival between the SDR fusion and uninterrupted KanR2 should only be observed in the presence of both kanamycin and zinc (Woods et al. 2020). Further, it is important to avoid certain additives in the medium that can interfere with the stress condition. In the case of M. smegmatis, these include the albumin and catalase within the ADC and OADC enrichment solutions.

Time considerations

Basic protocol 1: 2 weeks

Basic protocol 2: 3 days

Support protocol 1: 1–2 weeks