Molecular Cloning Guide

The Big Picture: Cloning into Zymomonas

Your goal is to:

Make a plasmid and then ultimately get that plasmid into zymomonas

What does that plasmid look like?

Let’s zoom into on what smaller steps those two big steps entail:

Two approaches to cloning


  • PCR amplify all fragments
  • Digest vector
  • Combine everything in 1 rxn
  • 1+ inserts
  • More flexible design
  • More difficult to troubleshoot
  • Design w/ NEBuilder


  • PCR amplify 1 fragment
  • Digest vector and fragments
  • Combine in 1 rxn. Repeat for next insert if needed.
  • 1 insert at a time
  • More limited design capability
  • Design in sequence viewer (e.g. SnapGene, Ape, CloneManager)

For both approaches, you’re going to prep a vector and insert(s) separately, then combine them.


Option 1: Gibson Assembly

Lots of info available on NEB’s website:

Gibson Assembly Reaction:

*Adjusted for molarity

To calculate amt of insert to add to the reaction:

150ng * (bp insert/bp vector) = ng insert to add

Here’s what’s happening in the tube:


Option 2: Traditional Cloning

Ligation Reaction:

Here’s what’s happening in the tube:

Screening for successful, correct plasmid construction

     We must ask: Did that work? Do I have the construct I want?


Construct Design

Start by drawing the construct you need

Remember to ask yourself:

  • Do you have appropriate markers?
  • What’s your plan for screening?
  • If you are expressing a gene from this plasmid:
    • Ribosome binding site (RBS)?
    • Promoter?
    • Start codon?
    • Stop codon?
    • Is it in-frame?
  • If you are using tags
    • N-terminal: Does your tag have a start codon? Did you remove the native start codon?
    • C-terminal: Did you remove the stop codon? Does your tag have a stop?
    • Is everything in-frame?

**If you answered “I don’t know” to any of the above… Talk to a labmate!**

Design a ribosome binding site (RBS)

  • If you are expressing a protein off your plasmid, your construct must include a RBS. Some vectors already have RBS+promoters ready-to-go, and others don’t. If you are using a vector that does not already have a RBS:
    • Go to
    • Create an account with your university email
    • Follow the steps to design an appropriate RBS for your situation (pics below)
    • Remember to add this sequence into your desired final construct

Designing Gibson Cloning

  • Contact Dave if you need a SnapGene license
  • Open your vector sequence. Make sure you’ve annotated the restriction enzyme cut sites. Open all relevant fragment sequences (genes, tags, RBSs, etc)
  • Go to
    • Click “+ NEW FRAGMENT” and input your complete vector sequence
    • After pasting your sequence, click “Process text”
    • Check the “Vector” and “Circular” boxes above the Parsed Sequence box (because the sequence you just pasted is your vector for cloning, and it’s a circular piece of DNA – not linear)
    • Name the fragment
    • Choose the method you will use to linearize the plasmid. You have two real options:
      • PCR: Amplify a portion of the plasmid using PCR, which creates linear pieces of DNA. If you don’t want to use the whole plasmid, you can amplify only the desired fragment.
      • Restriction digest: Cut open the plasmid with a restriction enzyme. You only need to cut the plasmid one time, to linearize it. If you want to use only a portion of the plasmid, you may cut multiple times to isolate your desired fragment.
      • If you are buying a synthesized vector (which is very unlikely, unless you are doing something really new and you happen to have a ginormous cloning budget…), check that box.
    • Add the sequences of your insert fragments using the “+ NEW FRAGMENT” button. Uncheck the Vector and Circular boxes (unless you are, in fact, using a circular DNA sequence as an insert; then you’ll have to choose a linearization method for that as well). Drag the fragments to get them in the right order. Make sure you input all genes in the correct orientation that you want for your final construct.

  • NEBuilder will output primers for you under “Required Oligonucleotides”
    • These are program suggestions – use your brain, too! Make sure these amplify what you want to amplify, are reasonable lengths, GC content, etc.
    • If you have any wiggle room on insert sequence, play with it to make optimal matching pairs of primers.
    • Everything is color coded to match it’s origin sequence
    • Primers are given as 5’-(overlap/spacer/ANNEAL)-3’. This means:
      • Sequence is written 5’ to 3’
      • 1st color = sequence acting as the adapter, creating overlap to the fragment of that color
      • 2nd color = sequence acting as a spacer (only present if you added a spacer)
      • 3rd color = sequence that this primer anneals to (and amplifies). Note that primers with the same ANNEAL color are primer pairs.
    • Validate your primers with IDT’s OligoAnalyzer
      • Login (No ordering on this account)
      • Ideal primers have:
        • 40-60% GC content
        • Low favorability for hairpin formation, dimerization (i.e. high ΔG)
        • Primer pairs have similar melt temps
      • Order your primers
        • Create excel document with primer names and sequences
Name Sequence
taggymctaggerson_F1 CGCTAGCTAGCTACA
taggymctaggerson_R1 CGATCGTAGCTAGCT
  • Record that you have a primer order in the ordering sheet on the lab website
  • Email/Teams your excel list of primers to Dave at

Designing Traditional Cloning

  • Contact Dave if you need a SnapGene license
  • Open your vector sequence. Make sure you’ve annotated the restriction enzyme cut sites. Open your fragment sequence
  • Linearize your plasmid
    • Choose a pair of restriction enzymes (w/ compatible buffers) that will cut your plasmid to open the space in which you want to insert a fragment. Compatibility chart available here:
    • Using two enzymes that leave different overhangs allows you to control which direction your fragment anneals into the open vector. If you only cut the plasmid once, the overhangs will be identical and your insert can anneal in either orientation, meaning you must do an additional screening step.
  • Identify the insert sequence you wish to add. You will need to design an upstream and downstream primer to amplify this sequence. Starting at the 5’ end and working with the sequence in the forward direction, choose ~14-20 bases at the start of this insert sequence.
    • If you have wiggle room about where your sequence starts, use the OligoAnalyzer (see Gibson section) to find a space with ideal primer characteristics:
      • 40-60% GC content
      • Low favorability for hairpin formation, dimerization (i.e. high ΔG)
      • Primer pairs have similar melt temps
    • Once you’re happy with this sequence choice, add the restriction enzyme recognition site to the 5’ end of the primer sequence.
    • Then, add an extra 4 bases, again at the 5’ end. This little tail can be any sequence – use it to balance GC content if needed. It’s purpose is to give the restriction enzyme some extra physical space to sit down on (since they are endonucleases), so the sequence is irrelevant. Your final primer will look like:
      • Where black is the 4bp tail, blue is the cut site, and red is the sequence of your fragment where the primer will bind and begin to amplify.
    • Repeat this process to design your reverse primer. I recommend setting your insert sequence to the reverse complement in SnapGene, and then acting in the “forward” direction while you design the primer, so that everything is consistently going in the correct direction.





Cloning CRISPRi Knockdown System into Zymomonas mobilis

(This was written utilizing protocols from the Guss, Pfleger and Dan Labs)




  1. Assemble CRISPRi plasmid with sgRNA of the desired gene to knockdown
  2. Transform CRISPRi plasmid into DH5alpha and AG4476
  3. Methylate CRISPRi and integrase plasmid
  4. Transform methylated CRISPRi plasmid and integrase plasmid into Zymomonas mobilis (AG4826)
  5. Verify integrated CRISPRi in the Zymomonas mobilis genome through PCR



– AG4476 (E. coli strain with Arabinose inducible Z. mobilis methylases) – this allows for pre-methylation of DNA to look like Z. mobilis drastically increasing the efficiency of transformation without needing a multi-deletion strain that has growth defects. All the plasmids you put into Z. mobilis get miniprepped from this strain.

– AG4826 (Z. mobilis ZM4::poly-attB cassette) This is a strain with poly-attBs inserted just downstream of the acetaldehyde dehydrogenase. Its exact position is between ZMO1753 and ZMO1754. This is the strain everything goes into.


– pGW39 (Non-replicative plasmid that transiently expresses the R4 Serine Integrase, which flips the cargo plasmid into the chromosome)

– pDC-Z095 and Derivatives (R4 attP; KanR; CRISPRi plasmid) This carries the IPTG inducible CRISPRi system, the guide can be swapped out via TypeIIs restriction. We use AarI (PaqCI is the isochiozomer from NEB).


Explanation of the System

ZM integrase system

AG4826 is the parent strain for most of our ZM-integrase work. This strain contains the 9X attB sites that match the attP sites in pJH204-pJH212.


Integrase system

The integrases catalyze recombination between their corresponding attB and attP sites. This is typically achieved by transformation of a strain containing a genomic attB (AG4826) with an attP/resistance marker-containing cargo plasmid (example – pJH204) and a plasmid which transiently expresses the integrase. Usually a suicide vector (example – pGW31). This results in recombination between the attB/attP sites, forming attL and attR which flank the cargo (example – AG4826 x pJH204).

Designing cargo plasmids for integration

We typically use either Bxb1 or R4 attP sites for integrations in Zymomonas. For selection markers, KanR (200 mg/L in ZM) or ChlorR (120 mg/L in ZM) work well for Zymomonas. We use an origin of replication that functions in E. coli, but not in Zymomonas. See pTM348 as an example. Swap out the resistance marker and attP site to create an orthogonal cargo plasmid for integrations into different attB sites.



For Zymomonas, we passage all plasmids (attP cargo, integrase suicide vectors, homologous recombination, ect.) through AG4476 + 1mM arabinose. This will methylate plasmids with ZM associated methylation pattern, improving transformation efficiency. Transformation of AG4476 can be achieved via typical electrocomp cell preps/protocols (E. coli Chemical Competency Cell Preparation in our protocols), followed by cPCR to confirm presence of the plasmid of interest.


Once confirmed, an LB + 1mM arabinose overnight culture will induce the ZM methylases, and a miniprep will yield ZM-methylated plasmid that is ready for transformation (aim for ~100 ng/uL DNA concentration).

Zymomonas mobilis competent cell prep


  1. Streak an ZRMG plate with AG4826 ((Z. mobilis ZM4::poly-attB cassette) This is a strain with poly-attBs inserted just downstream of the acetaldehyde dehydrogenase, its exact position is between ZMO1753 and ZMO1754) from -80˚C freezer stock. Incubate anaerobically for 24-48 hours at 30˚C.


  1. Use an individual colony to inoculate 50 ml RM/G2 media in 50 ml conical tube. Briefly vortex to mix. Remove 25 ml inoculated media and place into another 50 ml conical tube (giving 2 – 25 ml cultures). Place caps on loosely to allow gas to escape. Incubate in tube rack (without shaking) anaerobically at 30˚C overnight to OD6000 – 2.2.
    1. a. 50 ml culture will yield ~20 aliquots. Inoculate more or less depending on number of transformations.
    2. b. RM/G2 indicates 2% glucose. Recipe is as follows:
      • Per L: 2 g KH2PO4,  10 g yeast extract, 20 g glucose
      • For plates, use 15 g/L agar
      • Autoclave with “Liquid 30”


  1. Pellet cells. Use benchtop refrigerated centrifuge at 4˚C, max speed, 5-10 minutes.


  1. Resuspend each pellet in 25 ml ice cold 10% glycerol.


  1. Repeat steps 3 & 4 two more times (for 3 total washes).


  1. For the 4th and final wash, pool pellets together and resuspend in a total of 40 ml ice cold 10% glycerol. Then, pellet as above.


  1. Resuspend final pellet in remaining 10% glycerol after pouring off the supernatant. Bring final volume to 1000 µl.


  1. Make 50 µl aliquots. Use immediately or freeze at -80˚C.


Assembling CRISPRi Plasmid through Cloning

  1. Design your sgRNA and adding TAGT to the beginning of the forward (top) oligonucleotide and AAAC to the beginning of the reverse (bottom) primer oligonucleotide (You can order these from IDT, nothing special about these oligos)


  1. Set up your reaction to anneal your top and bottom oligonucleotide:



43uL ddH2O

1uL 100uM Top Oligonucleotide

1uL 100uM Bottom Oligonucleotide

5uL 10X NEB Cutsmart Buffer

50uL Total Reaction Volume


  1. Incubate at 95°C for 5min (strip tube in thermal cycler). Let oligos gradually anneal while cooling to room temp (remove from thermal cycler, leave on bench 20+ min). Use the annealed oligonucleotides to do a 20X dilution (5uL of the annealed reaction into 95uL of ddH2O) this will give you a concentration of 450ng/uL.


  1. Set up the assembly reaction to ligate the annealed oligonucleotides to the CRISPRi plasmid (You can use either of the two ratios of plasmid-to-oligonucleotides, 1:3 and 1:6, both work):


  1. Utilize a thermocycler to assemble your plasmid:


(37C for 1 minute, 16C for 1 minute) X 45 cycles, then incubate at 37C for 5 minutes and finally incubate at 60C for 5 minutes


  1. This will be your assembled plasmid; you can now transform into DH5Alpha and AG4476.


Transformation of DH5α or AG4476 E. coli


  1. Thaw aliquots of DH5α or AG4476 (prepared from coliChemical Competency Cell Preparation protocol) from -80°C on ice.


  1. Add DNA to DH5α or AG4476 tubes and flick.


  1. Let tube sit on ice for 30 minutes.


  1. Heat shock cultures at 42°C for 45 seconds.


  1. Leave on ice for 3-6 minutes.


  1. Add 900 µL of SOC (SOB + 2% glucose) media.


  1. Grow shaking for 1 hour at 37°C.


  1. Spin cells at full speed for 1 minute.


  1. Remove ~800-850 µL and resuspend in remaining volume (Around 120uL).


  1. Plate on prewarmed antibiotic plates and grow at 37°C overnight.


Methylate your CRISPRi and Integrase Plasmid


  1. To be able to transform a CRISPRi and integrase plasmid into Zymomonas mobilis (AG4826), both plasmids must be methylated. Streak each strain on an LB plate.


  1. Inoculate a single colony from each strain into LB + 1mM arabinose and grow aerobically shaking overnight at 37C.


  1. Miniprep the culture the next day, this will yield Zymomonas mobilis-methylated plasmid which is ready for transformation into Zymomonas mobilis (AG4826).





Transformation of Zymomonas using the integrase system


  1. Remember that all plasmids need to be methylated via strain AG4476 (with 1 mM arabinose). Place a 1mm electrocuvette on ice (one for each transformation), and thaw comp cells on ice. Add 100 ng of the methylated cargo (CRISPRi plasmid) and 1000 ng of the integrase (pGW39) to competent cells. The added DNA volume should be kept below 5 uL total to prevent arcing.


  1. Add to a 1 mm electroporation cuvette on ice.


  1. Electroporate at 1600 V, 200 Ohm, 25 µF. If arc occurs, try repreparation in comp cells diluted with cold 10% glycerol (1:1 ratio)


  1. Recover in 950 µl SOB + 2% glucose (Same media utilized in E. coli transformation) for 1 hour at 30˚C without shaking. It is okay to just use a 1.5 ml eppendorf tube. (You could also use 950 µl ZRMG instead of 950 µl SOB + 2% glucose but it is less efficient)


  1. Plate 100, 200, and 900 uL on ZRMG plates with appropriate antibiotics (Kan 200 mg/L). Note that only the cargo plasmid should be selected for, not the integrase expressing vector


  1. Incubate anaerobically at 30C for 48-72 hours
  2. Verify presence of plasmids with colony PCR. Primers can amplify an internal region of cargo plasmid or read across either or both of the attL and attR sites. This can be achieved by designing primers that read into the attB site on the AG4826 genome, and a set of primers reading into the attP site on the cargo. If properly spaced, 2 unique bands will be produced from the 4-primer PCR reaction if integration is successful. 1 band from the attB primer pair indicates no integration.
  3. Proceed with downstream testing/sequencing/comp cell prep – note that other att sites are still candidates for downstream integration