Protocols - Kortemme-Lab/wiki GitHub Wiki

Protocols on this page:

FPLC protocol

https://docs.google.com/document/d/1dn4rxjs7V2xk87Vihixf2tkpAPstD8nopLzRX8WUBpk/edit Requirements Sample: Approximately 1mg/mL in a suitable volume for the column 0.2uM filtered

Columns: Superdex 75 Increase 10/300 GL - small scale preparative (ug-mg) + analysis/characterization

Buffers: Filtered/degassed diH2O (~4.5 CVs) Filtered/degassed buffer (to run your sample in) (~2.5 CVs) Filtered/degassed 20% EtOH

Protocol

200pg 16/600 75pg 16/600 Superdex 75 increase 10/300 GL Column position A2 A4 Manually attach Maximum flow rate EtOH 20% 0.7 ml/min 0.7 ml/min 0.4 mL/min Maximum flow rate, water 1 ml/min 1 ml/min 0.8 mL/min Delta column pressure 0.3 MPa 0.3 MPa 1.8 MPa Pre column pressure 0.5 MPa 0.5 MPa 5.0 MPa Column volume 121 mL 121 mL 24 mL Loop volume Typically 5 mL position 2 Typically 5 mL position 2 500uL (loop position 1 - minimum 100uL)

Can check in Method Editor window. If your buffer is viscous (typically due to glycerol) use the flow rate for EtOH and watch the pressure

Prepare the pumps and column Wash the pumps with diH2O (not required to be filtered/degassed) Make sure line A1 is in the correct bottle. Manual: Pumps: pump wash A, inlet A1 Purge the pumps with a syringe All 4 pumps need to be purged. Open valve counterclockwise and wait for liquid to start flowing out. Attach syringe and withdraw around 5-10mL. Close valve and detach syringe. Wash the pumps with filtered/degassed diH2O Move line A1 into filtered/degassed water Manual: Pumps: pump wash A, inlet A1 If connecting a new column, wash the lines attaching to the column Connect spacer to the the lines you wish to use (e.g. 3A and 3B). Manual: Pumps: system flow 0.7mL/min, pressure control pre-column pressure Flow path: inlet A A1, column position (e.g. 3), flow direction down flow Timer (volume): 50mL Action: end Connect column Wash the column with filtered/degassed diH2O Start flow from manual: Pumps: system flow <flow for 20% EtOH at 4C> mL/min, pressure control pre-column pressure Flow path: Inlet A A1, column position (e.g. 3), flow direction down flow Monitors: autozero UV Alarms: delta column pressure , pre-column pressure Timer (volume): 2 column volumes Action: end Drip liquid into top of column until visible then screw in top connector Once liquid emerges from bottom of column screw in bottom connector Equilibrate the column with filtered/degassed buffer Move line A1 to buffer From manual: Pumps: system flow <flow for water at 4C> mL/min, pressure control pre-column pressure Flow path: Inlet A A1, column position (e.g. 3), flow direction down flow Monitors: autozero UV Alarms: delta column pressure , pre-column pressure Timer (volume): 1.5 column volumes Action: end

Inject the sample Wash the loop with 4X loop volume of diH2O → 0.5M NaOH → diH2O → buffer Close the syringe valve Manual: Flow path: injection valve position = inject → Execute Wash the connector with diH2O Rinse the entrance of the injection valve with a wash bottle Load the wash into the syringe (make sure there are no air bubbles) - use a new syringe for each wash and close the valve before each removal Slowly inject the sample into the injection valve (check that no air bubbles are flowing in W1) Loop valve: position Flow path: injection valve manual load → Execute

Filter the sample with a 0.2um filter Load the sample into the loop Make sure the injection valve is closed Flow path: injection valve position = inject Remove the syringe used for the last wash and the needle Load sample into a syringe with the needle and make sure there are no air bubbles With a wash bottle, put some water at the entrance of the injection valve Screw the syringe with the sample into the injection valve Push on the plunger to get rid of any air in the tip of the needle and check for bubbles again Inject the sample Manual: Loop valve: position Flow path: injection valve position = manual load Slowly inject sample into the loop Stop the system by pressing the stop button on the top toolbar Load wells into fraction collector View → Fraction collector to see current status Load by putting your well plate on the stand (should hear a click) then put onto the tray and slide the tray inside

Run the sample Run sample through the column Start buffer flow: Pumps: system flow <flow for water at 4C> mL/min, pressure control pre-column pressure Flow path: Inlet A A1, column position (e.g. 3), flow direction down flow, loop valve position Monitors: autozero UV, wavelength <value(s)> Alarms: delta column pressure , pre-column pressure Timer (volume): 1.5 column volumes, Action: end Optional: Fraction collector F9-C: Fractionation 96 deep well plate 2.00mL Inject sample once you see output to fraction collector or waste (fraction collector will let the first 0.5mL or so drip into waste before it starts collecting) Flow path: injection valve position = inject

Notes: If running multiple samples in the same buffer: Clean syringe adapter and inlet valve with water Make sure buffer level is sufficient Repeat sample loading and running steps If running multiple samples in different buffers: Equilibrate column in second buffer Wash the loop with 4X the loop volume in the second buffer Continue with normal sample loading steps

Clean and store Wash the column with filtered/degassed diH2O Move line A1 to diH2O filtered/degassed Manual: Pumps: system flow <flow for water at 4C> mL/min, pressure control pre-column pressure Flow path: Inlet A A1, column position (e.g. 3), flow direction down flow Monitors: autozero UV Alarms: delta column pressure , pre-column pressure Timer (volume): 2 column volumes Action: end Store the column in 20% ethanol Move line A1 to 20% ethanol filtered/degassed Manual: Pumps: system flow <flow for 20% EtOH at 4C> mL/min, pressure control pre-column pressure Flow path: Inlet A A1, column position (e.g. 3), flow direction down flow Monitors: autozero UV Alarms: delta column pressure , pre-column pressure Timer (volume): 1.5 column volumes Action: end

Collecting and processing NOESY and HCCH TOCSY experiments

https://docs.google.com/document/d/1JCD7oaHxNqL4n45FXPPUuiP8FLYVq_LNcj4WL52MpSY/edit To set up adiabatic decoupling: 13C decoupling: CPDPRG [2] = p5m4sp180.2 > then run pulsecal SPNAM 31 = Crp48,1.5,20.2 (should be automatic) P63 = 1500 us (should be automatic) SPW31 = 5.5638W (should be automatic) 15N decoupling: CPDPRG [3] = garp4.p62 > then run pulsecal P62 = 350 us (should be automatic) PLW16 = 2.4487W → need to manually set If you do pulsecal again it will reset PLW16 → need to manually set this again!

Data Acquisition On the 800MHz magnet: Create a new experiment with new (date_13C_15N_description_3D) Collect the following 2D experiments:

Experiment 1: N15 HSQC (FHSQCF3GPPH) - full spectral width In ACQUPARS, set: ZGOPTNS: -DLABEL_CN NS 2, TD F1 512 (may need to increase scans for larger proteins) Take note of: O1P (where the 1H pulse is centered) P1 (length of the 90 degree pulse)

Experiment 2: aliphatic centered CT C13 HSQC (HSQCCTETGPSP) - full spectral width Set O1P based on the value from expt 1 by typing o1p on the command line and entering the value manually Set P1 by typing getprosol 1H -10.62 on the command line In ACQUPARS, set: ZGOPTNS: -DLABEL_CN NS 4, TD F1 400 (can adjust TD as needed to modify time) O2P 43.000 Use 13C adiabatic decoupling: CPDPRG[2] p5m4sp180.2 → Run pulsecal. Should have set these automatically but check: SPNAM31 = Crp48,1.5,20.2 P63=1500us SPW31=5.5638W

Experiment 3: HH plane of the N15/C13 simultaneous NOESY Load NOESYHSQCGPSISMSP3D as the parameter set Set O1P, P1 as described above In ACQUPARS, set: TD F1 128, TD F2 1, TD F3 1024 Use 13C adiabatic decoupling as described above. Use 15N adiabatic decoupling: CPDPRG [3] = garp4.p62 → Run pulsecal. Should set this automatically but check: P62=350us. Manually set PLW16 to 2.4487W. If you run pulsecal again at any point after this, you will need to manually set PLW16 again. Suggested starting points: O2P 64.951 in30 0.0004110900 SW3 14.2022 SW2 39.1 SW1 11.8 Adjust SW of F1 to not cut off any NOEs

Experiment 4: HSQC plane of the N15/C13 simultaneous NOESY Copy the previous experiment with iexpno Set TD F1 to 1 and TD F2 to 128 Adjust SW F2 for 13C, in30 for N15, and center frequencies to make sure no signals have been cut off at the edge.

Experiment 5: HH plane of the HCCH TOCSY Load in parameter set HCCHDIGP3D Set O1P, P1 as described above. In ACQUPARS, set: TD F2 to 1 and TD F3 to 1024 SW1 to 7ppm Suggested starting points: O2P 25.662 SW3 14.2022 SW2 39.1

Collect the following 3D experiments:

Experiment 6: 3D N15/C13 simultaneous NOESY

Set O1P, P1 as described above. In ACQUPARS, set: TD F3 to 1024 and TD F1 to be as high as possible given time restraints (around 192)

Experiment 7: HCCH TOCSY

Set O1P, P1 as described above. In ACQUPARS, set: TD F3 to 1024, TD F2 to 96, and TD F1 dimension to be as high as possible given time restraints (around 192)

End with an N15 HSQC Before queueing everything: Signal present (run zg and stop once you see signal) Check DS/NS numbers Check O1P/P1 Check planes Check time - total should be around 120hr from when you started (including set up time) Turn on autoshim

HCCH TOCSY processing Same as the HcccoNH

NOESY processing In TopSpin drag your 3D experiment (PROCNO 1) into the display window In the processing parameters tab: Set SI for F1 and F2 to double what the corresponding TD values are in the acquisition parameters tab (ACQUPARS), BCFW, BC_mod, ME_mod, and SSB Reference the spectrum according to the parameters of the particular machine used Write a new process number with wrp (PROCNO 11) for 13C separately (will need to phase differently and do strip processing) Open PROCNO 11 - this will be the 13C side Project and phase the 23 plane as PROCNO 12 - see where the highest 1H shift is for the 13C HSQC side (e.g. around 7-8ppm) and strip process the spectrum Re-phase after strip processing the spectrum and save to 3D Phasing the 13 plane will be off by 90 degrees due to sensitivity enhancement (for the 3D use the phasing from the 23 plane) Fourier transform with ftnd 0 on the command line Open PROCNO 1 (the 15N side) Strip process the 15N side Re-reference the 15N NOESY - the F2 axis is incorrect (this experiment collected both 15N and 13C NOESYs simultaneously so the F2 axis is currently set for 13C) Go into the NOESY experiment folder and execute the sparky/getbruk script in this directory Note the SFO3 value (central frequency of the 15N dimension) and convert this to ppm ( - 81.0764602) / 81.0764602 * 10e6 117.105ppm Calculate the spectra width in Hz by taking the reciprocal of in30. Calculate this value in ppm as well by dividing by 81.0764602 (the 800 MHz DSS reference value) 2432.557347539 Hz 30.00325053ppm In UCSF format you define the upper and lower edges of the spectrum. Divide the SW in ppm by half and add it to the center frequency to get the range. Convert the 3rrr file to ucsf format. Go into terminal and convert the N15 NOESY: ./ucsfdata -a2 15N (change the nucleus name to N15 in the F2 dimension) ./ucsfdata -sw2 ./ucsfdata -f2 (changes the transmitter frequency) ./ucsfdata -o2 → will also update the top edge of the spectrum Compare to the HSQC

Collecting backbone and sidechain triple resonance experiments and data processing

https://docs.google.com/document/d/15v2rGB0ZvQcAlb3jDqvGGUpwhxVQeJCVapkXpJXjkoQ/edit?usp=sharing Experiment descriptions Avance 3D manual CBCAcoNH - This experiment, along with the CBCANH and N15 HSQC are needed for backbone assignment. Each strip contains the Ca and Cb signals of the i-1 residue.

CBCANH - Each strip contains 4 peaks - the Ca and Cb signals of the i and i-1 residue. The i-1 signals should be weaker (in some cases they won’t be visible).

CccoNH - This experiment is used for carbon side-chain assignment. Each strip contains peaks for all C atoms in the side chain of the i-1 residue

HcccoNH - This experiment is used for obtaining hydrogen side-chain assignments. Each strip contains the hydrogen shifts from the side-chain of the i-1 residue

  • These experiments are for proteins with 80-150 amino acids. For larger proteins, other experiments should be done with deuteration. The sensitivity of each experiment varies widely:

Sample requirements 300uM of N15/C13 labeled protein. Add 5-10% deuterated water.

Data acquisition See N15_HSQC document for information on loading your sample into the spinner and logging into the machine/opening TopSpin. Load your sample onto the 600ii magnet manually. Put your sample into the slot below the shuttle tube (you can manually move the red gate out of the way). Using the buttons on the left hand side of the machine, press the blue button to load. If your sample gets stuck in the birdhouse, gently squeeze the lever holding the sample to release it and press the emergency stop button (red) for 10 sec. Type new to create a new experiment - date_description_TR Turn on autoshim in the BSMS GUI at the bottom of the screen. This is important to maintain a good shim when your experiment runs for a long period (>4h). Collect the following 2D planes to test the experimental set-up and quality of the spectrum. Experiment 1: Standard N15 HSQC (full spectral width) Use this to determine which signals can be folded along the N15 dimension (any extreme/isolated peaks). Remember to set ZGOPTNS to -DLABEL_CN to decouple carbon and prevent splitting in the nitrogen dimension. By moving the center of the spectrum/changing the spectral width, peaks outside of the selected spectral boundaries will be “folded” to reduce the recording time while keeping spectral resolution constant.

Determine spectral boundaries that will decrease the N15 spectral width while not resulting in any overlapped peaks. You generally do not want peaks at the very edge of the boundaries. (center 117.65, width 19.5ppm, P1 9.66, o1p 4.699) If any peaks overlap, adjust the center frequency and spectral width until the aliased peaks no longer overlap with other peaks. Adjusting the center frequency won’t change the position of the aliased peaks relative to other peaks, but altering the spectral width will. Ideally you want the spectral width as small as possible (without having any peaks on the edges) to maximize resolution. Take note of the center N15 frequency by estimating the value using your cursor (the value of the N/H shift will appear in the top left hand corner).
Take note of the spectral width in the N15 dimension using the distance measurement tool to the right of the ↑ arrow (drag from the top to the bottom of the range and record the delta F1 value in the top left hand corner). Take note of P1 after pulse calibration by typing P1.

For example, for the following spectrum you may want to fold in the highest and lowest peaks marked in blue by changing the spectral width and center frequency to match the area of the spectrum delineated by the pink boundary.

Experiment 2: Folded N15 HSQC (reduced N15 spectral width) Use this to make sure folding did not result in overlapping peaks. Create another N15 HSQC experiment by typing iexpno. Type o3p - enter the new N15 center frequency Type sw - enter the new spectral width Type zg Enter multiple display mode by typing .md and overlay with the full spectral width N15 HSQC. Check to make sure peaks were folded to the right place. Once a desired center frequency/spectral width are identified, create a macro to automatically set these values by typing edmac (edit macro) > change source to user in the top right hand corner > type the following into the text box: getprosol 1H -7.99 o1p o3p 2 sw Save your macro with some name, such as TR_600

Experiment 3: CBCAcoNH - NH plane Create a new experiment by typing edc and select CBCACONHGPWG3D as the parameter set Type TR_600 Set the carbon dimension loop counter to 1: in ACQUPARS, set TD = 1 in the F1 dimension (squish all the signals into the NH plane) Set the number of scans to the minimum required by typing ns 4 Once the plane is done collecting, process it by typing xfb and store it in process number 2. Overlay with N15 HSQC from expt 2 to ensure everything looks well aligned.

Experiment 4: CBCAcoNH - CH plane Create a new experiment by typing iexpno Type TR_600 Set the nitrogen dimension loop counter to 1 and reset the carbon dimension loop counter: set TD = 1 for F2 and TD = 128 for F1. This plane shows all the CB/CA signals.

Experiment 5: CBCANH - CH plane (the NH plane will rarely have any signal so we don’t run this one) Create a new experiment by typing edc and select CBCANHGPWG3D as the parameter set Type TR_600 Set the nitrogen dimension loop counter to 1: in ACQUPARS, set TD = 1 in the F2 dimension Set the number of scans to the minimum required by typing ns 4 This spectrum will show Ca (pos) and Cb (neg) signals with opposite phases.

Experiment 6: HcccoNH - NH plane Create a new experiment by typing edc and select HCCCONHGPWG3D2 as the parameter set Type TR_600 Set the carbon loop counter to 1: in ACQUPARS set TD = 1 in the F1 dimension Set the number of scans to the minimum required by typing ns 16 Overlay this spectrum with expt 2.

Experiment 7: HcccoNH - HH plane Create a new experiment by typing iexpno Set spectral width to 7 (the data has been shifted) Type TR_600 Set the H loop counter to 1: in ACQUPARS set TD = 1 in the F2 dimension and TD = 128 in the F1 dimension Set the number of scans to the minimum required by typing ns 16

Experiment 8: CccoNH - NH plane Create a new experiment by typing edc and select HCCCONHGPWG3D3 as the parameter set Type TR_600 Set the carbon loop counter to 1: in ACQUPARS set TD = 1 in the F1 dimension Set the number of scans to the minimum required by typing ns 16 Overlay this spectrum with expt 2

Experiment 9: CccoNH - HCCCONHGPWG3D3 - run CH plane Create a new experiment by typing iexpno Type TR_600 Set the nitrogen loop counter to 1: in ACQUPARS set TD = 1 in the F2 dimension and TD = 128 in the F1 dimension Set the number of scans to the minimum required by typing ns 16

Before queuing everything, type ii (i.e. initialize interface - forces all the variables you’ve set to be updated and cycle through the pulse program to check things). Then for each queued experiment, type zg and see that you get signal then stop (just to make sure you don’t collect any empty experiments). Queue the following (will take approx. 40h): Experiment 10: CBCAcoNH CBCACONHGPWG3D TD F2 = 56, NS = 4 Experiment 11: N15 HSQC NS = 2 (diagnostic N15 HSQC to run in between each 3D experiment to monitor protein) Experiment 12: HcccoNH HCCCONHGPWG3D2 SW F1 = 7ppm, TD F2 = 56, NS = 16 Experiment 13: N15 HSQC Experiment 14: CccoNH - change F2 to 56 HCCCONHGPWG3D3 TD F2 = 56, NS = 16 Experiment 15: N15 HSQC Experiment 16: CBCANH CBCANHGPWG3D TD F2 = 56, NS = 8 - doubling the scans gives root(2) more signal Experiment 17: N15 HSQC Set up the aromatic experiments Experiment 18: HSQCETGPSISP2 - aromatic centered C13 HSQC (this is what we use to set up the aliasing for folding the NOESYs/TOCSY) Pulsecal to set the pulse power Modify 82.85ms in IN_F [usec], o1p: 120ppm (in the middle of the aromatics) in the acquisition parameters (ns 8, ds 32, td 1024 256) TD in F1 affects length of decoupling (too high w points may cause heating of the sample but it might be better to be consistent with the other experiments) D24 2h 22min total time

Experiment 19: Cb/Hd expt - HBCBCGCDHDGP Pulsecal to update all the pulses Set 1k of points in acquisition Use adiabatic coupling to reduce heating (CPDPRG 2 - p5m4sp180.2 - will change spw31 to ~ 2.8W) instead of garp (p63 - length of pulse, sp31 - shape/pwoer) Type getprosol 1H -7.something or pulsecal Pulses for a specific application have a number and power value Set D16 to 195us Do the minimum ns and 64 td in f1 at first Change to ns 360, 96 td in f1 (modify ns to change the total amount of time the expt is running) Experiment 20: Cb/He expt - HBCBCGCDCEHEGP Change CPDPRG 2 again Type pulsecal or getprosol to update parameters Change tD to 1k Theyre calculating delta6 - changed D16 to 195us (recovery after gradient pulse) - only complains when decoupling scheme is changed for some reason? Do the minimum ns and 64 td in f1 at first Expt 21: N15 HSQC Turn off autoshim.

Potential errors: Cannot automatically tune the probe atma exact → ATMA terminated with error: The ATM algorithm was not successful → type atma f1 exact (tunes proton) → if this doesn’t work type atmm (manual tuning) Tuning = side to side Click the < > buttons to move the V (at first nothing will happen and then it will jump due to the motor catching) Match = depth of the V Try to make the V centered about the red line and at maximum depth (should try to touch the X axis) Can start ATMA from this window if you want to Type wobb to look at the tuning Cannot lock onto the deuterium signal lock > Command ‘lock’ failed - Lock is OFF - try loading shim file in first (0_CURRENT) but if this doesn’t work eject the tube and ensure that the sample height is ok + open the lock window Can click the icon to the right of the stopwatch in the top menu to look at the lock/level panel > can increase lock power (-6.4) and gain (120) until you see the lock signal in the lock panel - can adjust the wheel for field to center the lock signal, DC = how far up and down the lock is Can try some other shim files as well Try shimming with the lock off with parameters lockoff ordmax=3 and right click sample icon to toggle lock off > try to get within 2-3Hz (can increase ordmax) May need to shim with another sample (sucrose) - run a 3D then do a tune (all the horizontal planes) then do a 1D > redo lock 3D gradient (put fast in the parameters to only do one iteration) Field drifts to -9999 > unknown cause but will cause data to collect and be unusable (full of noise) May be due to lock failing during the acquisition

Data processing In TopSpin drag your 3D experiment (proc. no. 1) into the display window In the processing parameters tab (PROCPARS): Set SI for F1 and F2 to double what the corresponding TD values are in the acquisition parameters tab (ACQUPARS)

BCFW 0.4, BC_mod qfil for F3 - filters a region out in the center of the spectrum that is 0.4ppm wide (water), set ME_mod to no (no linear prediction), SSB 2 in F3 (no undershoots) Reference the spectrum according to the parameters of the particular machine used:

Bruker DRX 500 Bruker 600ii Bruker 800 1H 499.9299507 599.9799391 800.1299314 13C 125.7071511 150.8646737 201.1922952 15N 50.6574610 60.7954380 81.0764602

Phase the acquisition dimension and then check the indirect dimensions For your 3D experiment, set the processing parameter TDeff to 1024 (half of TD in the acquisition parameters) to get better signal-to-noise. Protein signals tend to decay within 50-60 ms while the rest of the signal is mostly water/buffer:

2D: xfb → RAW 23 (N15 HSQC) plane number 1 in PROCNO 2 (only some things like the phasing will get written over to the 3D menu) Strip processing (in 3D) Look at range of the N15 HSQC with cursor (the value at the top will show the Hz index) - don’t cut off any peaks! in the acquisition parameter F3: STSR 128 (first output point of the strip spectrum), STSI 768 (total number of output points of the strip spectrum from the start) - these numbers should be multiples of powers of 2 (due to the data “block” widths being powers of 2 in Bruker) > don’t want things at the very edge of the spectrum and have most of the signals in the middle

Click save contours Enter manual phasing (.ph) Add peaks to phasing (should select peaks that are strong and well separated in the spectra - at least one in the middle of the spectrum)

Right click to set pivot point on the peak that is in the middle of the spectrum. Phase in the acquisition dimension first (rows in this case - F3). 0 order correction applies the same phase to all points in the spectrum whereas 1st order applies frequency-dependent phase correction. R will reset to the original. Click save. Phase in the indirect dimension in the same way. Click save 3D icon (will apply to the 3D). You can see the phase correction in the PROCPARS PHC0 and PHC1 values. Shouldn’t have really large values (e.g. should be around 100-200 in F3 and 1-10 in F2) or you’ll start to get “dishing” (waves in your spectrum) Return to the 3D and do the same phasing for the carbon (RAW 13 plane number 1). The H has already been phased (unless you’ve done sensitivity enhancement). Check the PROCPARS in the 3D to see how much the F1 PHC0/PHC1 changed by. ftnd 0 (process the data with the order defined in the pulseprogram 321) > stdout Click 31 > scroll through CH planes → will show peaks in 128 planes → find planes with phasing issues (e.g. really sharp peaks with tails) → r13 (use same PROCNO as plane no) this will read out the plane as a 2D To get complex data back: Advanced > special transforms → Hilbert transform in F1 (xht1) Go into phasing menu and do phasing for the C dimension. Store phasing into the 3D. Reprocess the 3D with ftnd 0. Scroll through the CH planes again. Click 23 > scroll through NH planes → will show peaks in 256 planes → find planes with phasing issues → r23 Repeat phasing process. Check to make sure carbon planes still look ok. PROCPARS 3D: SSB F3 2.6 (70 degree shifted QSINE bell) ME_mod: constant time in N/C dimensions: LPmifc for F2/F1 (linear prediction mirror image forward → forward prediction by increasing number of points used for prediction by mirror imaging backwards), NCOEF F2/F1 32, LPBIN F2/F1 num acquired points*2 (double the number of points), SI F2/F1 double ftnd 0 dlp Delayed linear prediction - basically what this does is no LPmifc in F2 at first and no appertization. Process proton dimension ft3; ft2 (process proton then nitrogen) → will see as you’re scrolling through 23 planes the peaks will look like HSQCs but peaks will change phase (modulated by the C frequency) → linear predict the carbon dimension first Now linear predict the F2 (LPmifc) → FIDs that we are FTing only contain a few signals so now the linear prediction gets more stable Calculate projections: Advanced > calculate projections > positive projection, plane orientation f1 f3, first plane 5 last plane 251, procno 999 --> will see things being well separated at this point Re-reference the N15 HSQC and overlay with your projection to make sure everything aligns nicely Will be the same for CBCANH (you will have phase of the peaks for Ca being opposite to the Cb) - do the first phasing on the CH plane (NH plane will have negative and positive signals but in the CH plane the negative peaks will generally be at the top and positive will be generally at the bottom) and CccoNH. For HcccoNH the indirect proton dimension has been cycled and needs to be processed differently. Do re-referencing as above but also add SWH/4 to the SR value (post-re-referencing) Compare the HH plane with the TOCSY HH plane - range should be about the same For the aromatics (Cd/Ce): Strip process to crop out aliphatic signals but otherwise process identically to an N15 HSQC

Overall: Spectrum Parameter set Linear pred SSB BCFW Aliasing Special instructions N15 HSQC fhsqcf3gpph None 2.5/2 0.4 None None C13 CT HSQC hsqcctetgpsisp None 3/3 0.05 None None CBCANH cbcanhgpwg3d LPmifc, LPfc 2.5/2/2 0.4 N15 Strip processed; reverse N15 TRUE CBCAcoNH cbcaconhgpwg3d LPmifc, LPfc 2.5/2/2 0.4 N15 Strip processed HcccoNH hccconhgpwg3d2 LPmifc, LPfc 2.5/2/2 0.4 N15 Strip processed, rescale (F1: SWH/4 - DSS) CccoNH hccconhgpwg3d3 LPmifc, LPfc 2.5/2/2 0.4 N15 Strip processed HCCH TOCSY hcchdigp3d None 2.5/3/2 0.05 N15 Rescaled Aro-centered C13 HSQC hsqcetgpsisp2 LPfc 2.5/2.5 0.05 C13 Strip processed Hd/Cb hbcbcgcdhdgp LPfc 2.5/2.5 0.2 None

He/Cb hbcbcgcdcehegp LPfc 2/2.5 0.2 None

N15 NOESY noesyhsqcgpsismsp3d None 2.7/2.5/2 0.4 None Strip processed; re-reference C13 NOESY noesyhsqcgpsismsp3d None 2.7/2.5/2 0.05 C13 Strip processed

Collecting a DOSY

https://docs.google.com/document/d/19v1iuY_EYbgRmv3t52egEbX5ErWuhRd_oUagP2dqGw4/edit Other DOSY guides: Oxford Wisconsin Columbia Topspin

Experimental background DOSY is diffusion ordered spectroscopy. It can be used to differentiate NMR signals from a mixture of molecules by differences in diffusion and can also be used to estimate the molecular size for a pure sample. In this experiment, the effects of diffusion are detected in a series of 1D experiments with varying gradient strength (i.e. reduction in signal intensity with increasing gradient strength is a function of the diffusion coefficient of the molecule(s)). This experiment typically takes 10-15 minutes to optimize the parameters and 5 minutes to collect the pseudo-2D experiment.

Sample requirements No isotopic labeling is required - we are collecting 1D 1H spectra. Approximately 100uM is sufficient. Complete temperature equilibration is necessary to prevent convection in the sample (>20 minutes). To attenuate convection, you can try keeping the sample volume to a minimum and using a 3mm tube.

Data acquisition Acquire a 1D 1H NMR spectrum Open a 0_1H_PS experiment to tune/lock/shim/calibrate the pulse/suppress water for your sample. See the N15 HSQC guide for more details. Create a new experiment with edc and select ZGGPWG (a simple watergate experiment) Calibrate the pulse with the O1P value from step 1. Set the following parameters: NS 8, DS 8, TD 1024, SI 2048, BCFW 0.4, qfil, D1=1s Type zg to acquire the spectrum Multiply the FID with em to improve resolution and S/N + remove artifacts Fourier transform with ft Phase then do linear prediction Inspect the spectrum. We ideally want to follow the intensity changes of shifted methyls in the 0-1ppm region that are well separated from the rest of the signals.

Optimize d20 and p30 Make a new experiment with edc (using the current parameters is fine) Set the following parameters in the ACQUPARS window: PULSEPROG = stebpgp1s191d GPNAM6 = SMSQ10.100 (setting gradient shape to smoothed rectangular) GPNAM7 = SMSQ10.100 GPZ1 = -20.00 GPZ6 = 1.00 (typical range 2-98) GPZ7 = -17.13 D19 = 0.0000624 TD = 2048 NS = 4 DS = 8 D6 = 0.6 P30 = 1ms (typical range 0.5 - 2ms) - do not set this too long or you will damage the probe! It should NEVER be higher than 3ms and P30/(D1+AQ) should NEVER be greater than 0.05 D20 = 100ms O1P determined during 1D 1H spectrum acquisition Run the experiment with zg to collect your reference spectrum (without attenuation) Process like the 1D 1H NMR spectrum and store in PROCNO 2 with wrp 2 In the same experiment, change GPZ6 to 95 and run the experiment again. Diffusion of your sample will attenuate the intensity of the spectrum. Process like the 1D 1H NMR spectrum Overlay this experiment PROCNO1 (95% gradient) with PROCNO2 (2% gradient) with the multiple display window .md Select the reference spectrum and scale it down to match the attenuated spectrum. The scaling factor should be 0.05-0.1. If it is not, you will need to optimize the parameters. If the scaling factor is >0.1 (not enough attenuation), you should increase d20 and/or increase p30. If you see no signal (too much attenuation), you should decrease d20 and/or decrease p30.

Run the DOSY experiment Copy the experiment with iexpno (use current parameters, no additional action) and change the PULSEPROG to the pseudo-2D version stebpgp1s19. ACQUPARS - symbol to the left of the double triangle - will open parmode window and you can change acquisition dimension of dataset from 1D to 2D. TD F1 32 (number of steps), NS 8, DS 32, FnMODE for F1 QF Run an AU called dosy on the command line First gradient amplitude = 2 Final gradient amplitude = 98 Number of points = 32 Ramp type: l (linear) Start acquisition Change the processing parameters to those typical for an N15 HSQC Apply zero-filling in the F1 dimension (SI in F1 = 2* TD in F1), etc. Process with xf2 and open the multiple display window with .md. Click this icon and hover over the shifted methyl region to see slices (scroll to change intensity). The slices will indicate if you need to further optimize d20/p30:

Analysis Processing Read the first FID by typing rser 1 and process it by typing efp then absn. Correct the phase by running the apk command (and doing manual phasing with .ph). While in the manual phasing mode, press the nD save button to apply the phasing to the other 1D spectra in the series. In the 2D mode, type xf2 to process and phase the dataset in the F2 dimension Set ABSF1 and ABSF2 to 1000.0 and -1000.0, respectively. Execute abs2 to baseline correct. Type setdiffparm to transfer important parameters to the DOSY module in TopSpin. Type rser 1 to bring up the 1D 1H NMR with the highest intensity From Analysis, open the T1T2 module from Dynamics Click Peaks/Ranges and select manual integration Integrate some shifted methyls in the low ppm range Click save as → export to relaxation module Click the Relaxation module from the T1T2 toolbar Click the > icon and select Intensity to fit the data (should appear like the curve above - exponential decay)

Estimating molecular weight https://pubs.acs.org/doi/pdf/10.1021/acs.jpcb.2c03554 You can estimate the molecular weight with the Stokes-Einstein Gierer-Wirtz estimation (Excel sheet: https://nmr.chemistry.manchester.ac.uk/?q=node/432)

If your Topspin is not compiling AUs correctly (happens with Macs frequently), try the following: Install the Command Line Tools package, open a terminal and type xcode-select --install then follow the prompts. In /opt/topspin4.1.1/exp/stan/nmr/au/makeau: Uncomment line 20 (i.e. remove the # from # $opt_native = 1; Remove -Wl,-lcrt1.o from line 494 (so it reads 'MAC_INTEL' => ' -Wl,-w'.) In /opt/topspin4.1.1/prog/include/lib/libcb.h: change #include <values.h> to #include <limits.h> on line 28 Beneath this line add a line reading #define MININT INT_MIN And also add a line reading #define MAXINT INT_MAX In /opt/topspin4.1.1/prog/include/lib/uni.h: Comment out line 182 (i.e. change attribute ((format (printf, 1, 2))) to //attribute ((format (printf, 1, 2)))

Acquiring an N15 HSQC

https://docs.google.com/document/d/1CoQb7r7V4nWjDKA2qgIaKpCEPdQAWPMtXL5KX9LxkoU/edit

Topspin tips: https://docs.google.com/document/d/1ksYcggsa5NLCiGOM4MsCZ56hyQmYHBHEIFjXbRVMwPE/edit?usp=sharing

Experiment description The N15 HSQC is like a fingerprint for your protein and is typically the first heteronuclear expt performed. It shows H-N correlations (mostly backbone amide groups but also Trp, Asn, and Gln sidechain groups). It is a good primary assessment of whether your protein is stably folded, has dynamics, or binds to ligand.

Sample requirements At least 50uM of N15-labeled protein (final volume of 500uL in a standard 6mm NMR tube, 250uL in 5mm shigemi, 170uL in slotted shigemi). Ideally >100uM of protein. If you need to add reagents (e.g. for a titration series), use a standard 6mm NMR tube. If you will not need to transfer anything into your sample and getting a high protein concentration is an issue, use a shigemi tube. To add reagent to your NMR tube, add a small amount of your reagent via regular pipette to the side of the tube. Using a long glass NMR pipette, transfer your sample over the area you added the reagent several times, taking care not to introduce bubbles. Link for how to transfer your sample into an NMR tube

5-10% deuterated water (D2O) Add this before you transfer your sample into an NMR tube!

Data acquisition Log into the desktop at the NMR machine > Open TopSpin from the Applications dropdown menu Check to make sure the temperature is suitable (lower menu bar in the TopSpin window) Load your sample into the magnet For standard 6mm NMR tube/regular shigemi: Insert tube into spinner (blue) Insert tube/spinner together into sample depth gauge (clear plastic) and center the sample volume to the center of the coil marked (don’t just push all the way down)

Insert spinner/tube into the sample exchanger carousel Ensure no sample is currently loaded in the magnet Type sx into the commandline to inject your sample For a salt tolerant susceptibility matched slotted shigemi tube: Insert tube into slotted spinner Remove the carousel Type ej to start air flow Add your sample Type ij to inject sample Read in the default shim file by typing rsh and selecting 0_CURRENT Create a new experiment by typing new NAME: date_description_initials EXPNO: 1 Directory: your home data directory (i.e. /home/data/username/data) Read parameter set: FHSQCF3GPPH (fast HSQC) Set solvent to H2O+D2O [X] Execute getprosol (will load calibrations for this particular magnet) If your sample is C13 labeled as well, go to the ACQUPARS tab > click the step function icon > search ZG > type -DLABEL_CN into ZGOPTNS (decouples the carbon/nitrogen nuclei) Tune the probe by typing atma exact. This will tune the coil to the required frequency and match the coil to the correct impedance. Lock the sample by typing lock and selecting H2O + D2O. This will keep the field stable (the spectrometer will correct field drift as it occurs and the drift is measured using the frequency of your solvent’s deuterium resonance). Shim your sample. You need a homogenous magnetic field over the entire sample volume in the probe’s detection coil or else you will have poor resolution/sensitivity. Shimming adjusts the field’s homogeneity by applying different electrical currents to coils (shims) located at strategic places around the sample in the magnet. Open the TopShim GUI in the upper toolbar Dimension [X] 1D Optimization: Solvent’s default Optimize for: 1H Parameters: If using a standard 6mm NMR tube, type ordmax=8 (this will allow higher order shims to be used to correct for the ends of the sample). If you receive the warning ‘BSMS limits exceeded’ that means the software tried to adjust one of the shims to the absolute maximum of its range. You should in this case set ordmax=5 to optimize the lower order shims first and increase to ordmax=8 later. If using a shigemi tube, type shigemi. Start. This will tune in the Z direction (where most of the heterogeneity occurs). Your final B0 stdDev should be under 2-3Hz (but ~0.1Hz is best - you probably will not be able to get down to this level with a shigemi tube). Go to the Report tab to monitor the shim If the shim is not good (>3Hz), try using automatic gradient shimming by setting before [off], after [Z-X-Y-XZ-YZ-Z], only [X] to shim in the horizontal direction and then repeat shimming in the Z direction again (return to the original shim settings). Important: If during the process of shimming (which can be long and tedious), you get a good shim of around 1-2Hz, write this out to a temporary file by typing wsh in case you are not able to get a better shim. To suppress the water signal and calibrate the 1H pulse: Drag data/0_1H_PS_NEO/1 into the display window (you may need to scp this into your personal home directory first from the nmrsuser super user directory) and type pulsecal. Type gs and close the warning window about high intensity. Click the Offset tab in the GUI that opens > click the icon to the left of STOP to execute real time Fourier Transform Can also type rga then zg on expt 1 > look for the O1 hz value (around 3600) and then type that in for O1 then restart the calibration at that value with a sensitivity of 1 (FID area should decrease) Find out which power is for water suppression for HSQC Move the slider to the optimum frequency to suppress the water signal at 4.7ppm (Note: to move the slider, just click once above or below where the slider currently is and wait for the spectrum to adjust. Do not click and drag.) Click save and type o1p into the commandline to see what the 1H frequency is now centered at (should still be very close to 4.7ppm) Click 2D in the upper left toolbar to reopen your HSQC experiment Type o1p and enter the frequency you determined from (d) Calibrate the 1H pulse by typing pulsecal Set the number of scans. Start with 4 by typing ns 4. This will take about 40 minutes to run. If you have a lower concentration of protein (~50uM), you may want to increase the number of scans to 8. Start the acquisition by typing zg. Monitor periodically by processing the data as the acquisition is running. Fourier transform your data by typing xfb. Phase your data automatically by typing apk2d. If this is not sufficient (i.e. you are still seeing large negative signal in your spectrum), manually phase by selecting the Process tab at the top of the window > adjust phase > place crosshairs at several peaks with strong negative/positive signal at distant regions of the spectra > click and hold 0 while dragging up and down to phase with 0-order correction > repeat with 1st order correction if needed. If you are running multiple conditions for one sample, type iexpno once the acquisition is finished to copy the experiment parameters to the new condition and create a new subfolder in your main experiment directory. If you want to run a completely new sample, type new. If you remove your sample from the magnet, you will want to tune, lock, shim, and suppress the water signal/calibrate the proton pulse again. Eject your sample by typing sx ej. Log out when finished (this determines the pricing!)

Clean your tube! To clean the NMR tube, remove your sample and transfer it to an eppendorf tube with the long glass NMR pipette. Using the vacuum apparatus (basically a long plastic tube in an erlenmeyer flask), thread the plastic tube into your NMR tube. Dip the other side of the tube (with the adaptor attached) into cleaning solvent and firmly press straight down to seal the tube with the flask stopper. Do not bend or else you can break the tube. Do this for water 3X (getting the water into the bottom of the tube where your sample was), ethanol 3X, and acetone 2X. On the last acetone wash, keep the tube sealed for a minute to dry the residual acetone from inside the tube. If the sample was very old, store in HNO3 overnight and rinse with copious amounts of water the next day with a final acetone rinse. Clean cap by storing in ethanol. Let tubes dry upside down onto a kimwipe layer in an open Falcon tube at least overnight before using.

Amy’s notes If you have a good shim, you’re basically home free. If you cannot get a good shim, you could be stuck there for a long time. Try to eject your sample and re-adjust in the sample depth gauge especially if you have a very small volume of liquid. Your shims will NOT always get better the more you shim (sometimes you will jump from 0.8Hz to 5Hz). If things are looking bad and not getting better, restart from 0_CURRENT. If you really can’t get a good shim, for proteins we really just care about the water suppression. Do the pulsecal/gs step and see what the water suppression looks like (bad suppression would look like a giant antiphase peak that never gets smaller) - if it looks ok, proceed even if the shim is bad (e.g. around 3Hz).

For every 2X increase in concentration, the acquisition time decreases by 4X. It is best to have a sample that has a concentration >100uM.

If lock drifts to the top type “loopadj”

To reset the carousel (light red): Press the button on the front for 8 seconds to shut the carousel off CAREFULLY remove the carousel - make sure your tube is NOT in between the bore and the carousel (can tell if you stick your finger down into the sample holder or shine a flashlight inside). Lift the carousel slightly then slide it off slowly to prevent snapping your tube if it is partially still in the carousel. Place carousel aside and press the front button once to restart. Once the light at the top turns white, the carousel is ready to be replaced (there is only one configuration that will “click” into place so that the exchanger knows which position is #1).

To manually load samples on the 600ii: Don’t do this without talking to Mark first.

Unplugg voltage near compressor and turned blue adapter 90 deg at the back of the magnet. Type HA in commandline to open ethernet addresses. Click open next to the BSMS menu and clicked service link. Pass is 1964 and user is service. Go back to main menu. Click sample handling then sample transporter control. Change lift mode from Sample Transporter to standard bsms lift and hit Set. Refresh to make sure it’s set.

Turn valve back first, plug back in power block, then switch transport control back also. If there is a red light it should go green again - you can type ii to initialize interface (any settings set on workstation will be forced on the hardware). Flashing red - going back to starting positions. May want to cycle power (pull out plug wait 10s and put it back in)p

For variable T series: Re-shim and re-tune at each T and redo water suppression/pulsecal Heater power should be non-zero to have temperature control (4-5%?) Set power to maximum on the chiller to get down to 5C Set power back to medium once you’re at 15C?

Manually checking NMRtist assignments in CCPNMR

https://docs.google.com/document/d/1rCJr6NRI8JGEfTvRHyufCKa5Vr6ZBX1W_VMqPcyP1Hg/edit Purpose: You have collected NMR data, processed it, and run it through NMRtist to obtain a structure and/or chemical shift assignments. The machine learning algorithm can get things wrong at times + sometimes there is a flexible region of your protein that complicates automated analysis. Manually inspecting your data is required for accurate assignment/structure calculations.

Required software: CCPNMR

Copy and paste your chemical shift list from NMRtist (ARTINA_structure_calculation/1st_structure_proposal/structure.prot) into a text editor. This file contains the chemical shift assignments of each atom in your protein (except some NMR silent atoms). See this full guide for atom naming conventions (note: when there are chemically equivalent hydrogen atoms, these will often be denoted by a pseudo-atom name such as QB or HB% but when they are chemically distinguishable they will be called HB2 and HB3, for example)

Open CCPNMR and load in your NMR datasets: 1H,15N HSQC 1H,13C HSQC 15N/13C NOESYs Backbone assignment 3D spectra (e.g. CBCAcoNH/CBCANH) Sidechain assignment 3D spectra (e.g. HcccoNH/CccoNH) HCCH TOCSY - rotate the axes by typing xy Aromatic experiments (CBHd/CBHe)

In CCPNMR, drag the following spectra onto your screen in separate panels: HCCH TOCSY HcccoNH CccoNH

I recommend you check all the assignments but you can also focus on a specific region. You should pick a solid “starting point” in your sequence where you can fairly confidently identify that the assignments given by NMRtist are correct and work sequentially from there. For example, residues with characteristic proton assignments are ALA, ILE, GLY, LEU, SER, THR, VAL. This website is quite useful for referencing chemical shifts. Find a stretch in your protein that has 2+ of these residues consecutively (e.g. …QEISQN…). Depending on where your desired region is relative to this characteristic, you will go in either the i+1 or i-1 direction from here. For your first pass, I would ignore aromatic assignments and methionine methyl group assignments.

Imagine your problematic region is in the i-1 direction compared to your characteristic region:

Look at the assignment for N and Hn for the residue at the i+1 position relative to your characteristic region (e.g. …QEISQN…). Navigate to this “strip” in the HcccoNH/CccoNH (These are 3D experiments where each “plane” has the same N shift and the X-axis corresponds to the Hn shift. The Y-axis is the Hc or C shift of the aliphatic atoms in your sidechain. If you navigate to the Hn/N strip of residue position i, you will see the Hc/C shifts of the sidechain atoms for the i-1 residue position).

The peaks at this strip should be fairly easy to match to an amino acid identity (reference the guide above for reference chemical shifts). If they are consistent with the expected amino acid, check that the assigned shifts from NMRtist correspond to what you observe in the spectra. Navigate to the N/Hn strip for the characteristic residue closer to the C-terminus (e.g. …QEISQN…). This should have peaks consistent with the i-1 characteristic residue (e.g. …QEISQN). Check that the peaks align with the assignments from NMRtist. If everything looks consistent, this is now a solid starting point for checking the other assignments sequentially.

Proceed similarly in the i-1 direction. Specifically, Navigate to the N/Hn strip of residue position i in the HcccoNH/CccoNH. Briefly check that the pattern of peaks is consistent with the amino acid in the i-1 position. Look at the sidechain carbon assignments from NMRtist. Open multiple HCCH TOCSY panels with the “+” button in the top toolbar (as many panels as there are carbons in the sidechain with assignments). Navigate to the plane corresponding to one of the sidechain carbons for each panel (e.g. panel 1 is CA, panel 2 is CB, panel 3 is CG, etc.). Verify the Hc/C shifts. The HCCH TOCSY is a 3D experiment where each plane has the same C shift, the X-axis corresponds to the shift of the Hc atom connected directly to the carbon, and the Y-axis corresponds to the shift of all other Hc atoms in the sidechain (the peak on the diagonal is the Hc atom directly connected to the carbon). If it is too difficult to verify due to overlap in the HCCH TOCSY, you can take a look in the C13 NOESY as well. If the amino acid has an amide group (e.g. Q/N) then check the amide assignments using the N15 NOESY. If you notice any inconsistencies between the NMRtist assignments and the spectra, modify the chemical shift list accordingly.

Once you are done your first pass, go back to look at the aromatic assignments and methionine methyls. Open the CBHe/CBHd, 13C NOESY, and 1H,13C-HSQC spectra To verify aromatic assignments (after verifying the aliphatic assignments), I usually do the following: Look in the CBHe/CBHd spectra (overlaid) at the peaks corresponding to the CB shift of interest. Hopefully these are well separated from other peaks and the Hd/He frequencies can be easily verified from these spectra. Look in the 13C NOESY in the aromatic region (C shift 110-150ppm) for strips of peaks that correspond to what you would expect for that residue to assign the C shift (i.e. you see peaks corresponding to HA/HB of that aromatic residue and peaks corresponding to the shift of other H atoms close in spatial distance to the aromatic H atom in question). It may help to mark the Hb/He/Hd shift (if known) and scroll through C planes to find the strip of interest. Many times the Hz strip (of PHE) will overlap with the Hd/He strip and it will be hard or impossible to distinguish them. In that case, I delete the Hz assignment from the chemical shift list. To verify the methionine methyl assignments, open the 1H,13C HSQC. Look in the region of C 12-22ppm and H ~2.5ppm. There will be some quite intense signals there in your 1H,13C-HSQC arising from methionine methyls. Navigate to the strips in the 13C NOESY corresponding to these signals and try your best to match them to the methionines in your sequence. This can be extremely difficult since the methionine methyl signals are often filled with artifacts. If you aren’t sure, leave that shift out of the chemical shift list.

Protein expression for labeled proteins-NMR

https://docs.google.com/document/d/1zEbQfxdu9-VKnXdWMNAfiWw0nTHf_24uPJ8AVPD3ECg/edit?usp=sharing Expression Materials To make 1L of M9 media: 100mL of 10X M9 salts 20mL of 20% glucose

  • If making 13C labeled media, replace this with 4g/L of U-13C-glucose 1mL of 1M MgSO4 1mL of 1000X vitamins 10mL of 100X trace elements 1mL of 1000X kanamycin Water to 1L 0.3mL of 1M CaCl2 (add dropwise and swirl vigorously - will form insoluble CaSO4 at first)

Stock solutions: M9 stock solution (10X) Na2HPO4 (60g/L) KH2PO4 (30g/L) NaCl (5g/L) 15NH4Cl (5g/L) Dissolve in 800mL water, adjust to pH 7.2 with NaOH, add water to final volume of 1L and filter.

Vitamins (1000X) Dissolve 29.5mg in 25mL water. Add drops of 1N NaOH until biotin is dissolved. Dissolve 29.6mg thiamin-HCl. Add water to 29.5mL. Filter. Can combine with thiamin.

Trace elements (100X) Reagent Final conc Stock conc Vol. for 200mL stock EDTA 13.4mM 500mM 5.36mL FeCl3 3.1mM 1M 620uL ZnCl2 0.62mM 1M 124uL CuCl2 76uM 0.1M 152uL CoCl2 42uM 0.2M 42uL H3BO3 162uM 1M 32.4uL MnCl2 8.1uM 1M 1.62uL

Mix all components and filter. Keep this in a container protected from light.

Suggested culture volumes: For 15N HSQC, try 100mL total final culture volume For structure determination, try 500mL total final culture volume

Protocol Freshly transform BL21 with design Make a 10mL M9 media starter culture and inoculate a single colony for approximately 16h Dilute into final culture volume at an initial OD 600 of 0.05 Induce protein expression at an OD 600 of 0.6-0.8 with an appropriate amount of IPTG Culture at predetermined temperature for desired amount of time

Purification Materials Equilibration buffer (50mM Tris pH=7.5, 300mM NaCl, 10 mM imidazole) Wash buffer (50mM Tris pH=7.5, 300mM NaCl, 25mM imidazole) Elution buffer (50mM Tris pH=7.5, 300mM NaCl, 250mM imidazole) - not needed if doing enzymatic cleavage Enzyme cleavage buffer - depends on your enzyme Ni-NTA resin Enzyme capture kit (ideal, but not needed if sample is only needed for the next few days) Protein storage buffer - depends on downstream applications

Protocol Spin down cell pellet (8000g, 10min) and optionally freeze at -80C Lyse the cells with sonication or detergent and add a protease inhibitor cocktail pill. Spin down lysate 18000rpm, 30min While lysate is spinning down, prep your Ni-NTA resin. Add 2mL Ni-NTA resin slurry per 1L culture to a centrifuge-compatible tube Add at least 2 resin bed volumes (aka half of the slurry volume) milliQ water and spin down 750 rpm 2-3min + decant to remove EtOH (decant either by carefully pouring out supernatant or pipetting) Add at least 2 resin bed volumes of equilibration buffer, resuspend, and centrifuge + decant Mix lysate supernatant with an equal volume of equilibration buffer Pour supernatant onto beads and let proteins bind - 30 min-1 hour on a nutator at 4C ideally Wash pellet 3X in 2 resin bed volumes of wash buffer (centrifuge 750g, 2-3min) Wash pellet 1X in 2 resin bed volumes of enzymatic cleavage buffer (centrifuge 750g, 2-3min) Follow instructions from enzyme kit to cleave the His-tag off of your protein. Notes: Do a small scale test digest with 10ug of protein immobilized on the Ni-NTA beads and separately 10ug of protein in solution. For each condition (4x, 2x, 1x, 0.5x units of enzyme) take an aliquot at 2h, 4h, 8h, 20h. In general, more thrombin will be required to cleave directly off the beads. Prepare a fresh mL of Ni-NTA resin (wash with water, protein storage buffer) for the reverse column. Centrifuge cleaved protein solution and fresh resin separately Repeat 3X: Add supernatant from the cleavage reaction onto fresh Ni-NTA resin and invert to mix. Centrifuge and transfer to a fresh tube. Add 1 resin bed volume of protein storage buffer onto cleavage resin to increase yield Centrifuge “new” resin 2500 rpm, 5min and transfer the supernatant to a fresh tube If using a cleavage capture kit, mix with cleavage enzyme immobilizing resin now. Filter all of the collected supernatant from the fresh tube with a syringe into another clean tube Concentrate/buffer exchange into your protein storage buffer with an appropriate MWCO Amicon column or dialysis. Depending on how many spin downs you do, this may take a couple hours or o/n for dialysis.

Running CD

https://docs.google.com/document/d/1Ua_YpuHmj_iOqq-3MDT5Ph5fY1zvvjRTEQYJrpRhTUk/edit Introduction CD spectroscopy is used to determine the secondary structure of molecules. CD (measured in molar ellipticity) is the difference in absorption of left-handed and right-handed circularly polarized light and can be observed in molecules with chiral centers (e.g. proteins). Tm can be measured by following changes in molar ellipticity with increasing temperature. You can also use it to monitor structural changes, conformational stability, etc. It is label free and non-destructive.

Instrument We have a JASCO J-710 CD spectropolarimeter with a Peltier temperature controller and SpectraManager software for data collection and analysis (not supported by JASCO any longer).

Sample requirements Pure solvent - this will be your blank. Should be the same solvent that your protein is dissolved in. Take care that the concentration used does not cause the voltage to surpass 700V at any point during your scan. Typically, for KCl, I use 10mM. Protein sample(s) - you typically want enough protein to cause a minimum in your CD spectra around -20 mdeg (do not go below -50 mdeg). Any higher and you risk surpassing the 700V limit. The signal is linearly proportional to your protein concentration, so adjust accordingly. For a 10kDa alpha helical protein, I typically use 2-3uM of protein for a 1mm cuvette (you may need 2-4x more for a beta-sheet protein). The total volume should be around 500uL for a 1mm cuvette. Make sure the nitrogen tank has enough for your run! If less than half full, monitor the tank as you are running your experiment. Never run an experiment without nitrogen gas.

Prepare your sample

Add your protein using a pipette to a cuvette (these are in the left drawer under our UV-Vis spectrophotometer). Do not have bubbles in the cell. Insert the cuvette into a black metal holder. Be consistent with how you load your cuvette and how you place it in the machine, e.g. keep clip tension pushing towards the right with cuvette text facing the right

Prepare the machine Check the “fuel gauge” on the liquid N2 cylinder to make sure the tank is not empty. If it is, order another N2 cylinder. Make sure the regulator valve is turned opened slightly Turn the valve that controls flow from the cylinder into the regulator to “open” as well Use the regulator valve to fine tune the pressure to approximately 20 PSI Flip left-hand switch on the silver flow adapter box (behind the monitor) and adjust the flow rate with the dial to 15L/min (should be pre-set) Press red square button on the adapter box to illuminate the “LINE” light on the power supply box beneath the instrument Let the system purge with N2 for 10 min Check behind the small door on the CD machine (press to open) and confirm that the balance meter mode is set to PMV and the HT switch is set to AUTO On the Jasco Power Supply, turn on lamp (left, red button) and wait 15 seconds (should hear slight whirring as the fan turns on) Turn on the photomultiplier (right, green button) and let the lamp equilibrate for 30 min. You should never have the green switch flipped on without the red switch also being on. Note: If the green light does not turn on, follow the powering off instructions below and try again in another 10 minutes. Turn on the temperature control box (beside the monitor) and temperature bath controller (beneath the table) Open Spectrum Manager Upon startup, the instrument checks various levels. If the amplitude is listed as “too low” but the other settings are fine, hit “Ignore” to continue. If the lamp is old, HT may also be “too high”, but this only happens at very low wavelengths, so you can ignore if this happens. Select Spectrum Management If using the temperature controller, connect to it using Measurement > Accessory and select “PELTIER” Adjust temperature on the box to the left of the monitor by hitting “Stop” > “Enter” > adjust using arrows > “Start” Using the Control > Move wavelength option, set the wavelength and record the voltage in the notebook in front of the computer at the following values, in order: 600nm, 190nm, 253.7nm. These values should be similar to the last time the CD machine was used.

Load sample and ensure concentration is appropriate The voltage should never exceed 700V - you will damage the photomultiplier! Load your cuvette into the machine with the text on the cuvette facing the monitor (i.e. the wide side of the cuvette should be perpendicular to the longest length of the machine) Do a test scan to check that the voltage will not go out of range: Measurement > Parameters > Start: 230nm, End: 200nm. Click “Start” and hit “Yes” when the dialogue box pops up. Make sure the voltage is not projected to exceed 700V - if it looks like it will, hit “Stop”! Adjust your protein concentration accordingly.

Collect data CD spectrum From Spectrum Manager, select Spectra Management Set parameters:

Tab Setting Notes Value Parameters Sensitivity Keep at standard for proteins Standard Start/End Far-UV range for protein secondary structure 200-280nm Data Pitch Number of points taken during scan - doesn’t matter as much for continuous mode 0.1nm - 0.5nm Scanning mode Leave on continuous (faster and just as accurate as step) unless you for some reason need discrete wavelength data Continuous Scanning speed Rate at which CONITNUOUS mode collects data 100nm/min Response Determined by scanning speed 1s Bandwidth Can increase to 2nm if signal is low (especially for beta proteins) 1nm Accumulation Can do 3 for blank (this is the number of scans to average together) 5 Data mode Channels The data to be collected CD and HT Data File Auto save Whether to automatically save your data at the end of a run Keep checked and choose file name/directory Option Various Just notes for the user - record anything you want to about your sample

Run your buffer blank (will be used to subtract the baseline later) Dump out buffer carefully then load your sample into cuvettes, load into cuvette holder, place in machine (use the same cuvette that held your buffer - don’t dry) Before running, update the parameters (especially the file name so you don’t overwrite your blank) In Spectra Analysis, Processing > Subtraction > select buffer curve then save as a txt file

Thermal melt Recommended: perform a CD spectral scan before starting a variable temperature measurement From Spectrum Manager, select Variable Temperature Set parameters:

Setting Notes Value Wavelength Wavelength to monitor over the course of T scan

Start/End Start and end temperatures Typically 25C-95C Data Pitch Not sure if this matters? 0.2C Delay time Time to wait before measurement after target temp reached 60s Temperature slope How fast the temperature increases 1C/min Options If you want to do a reverse scan (with the same temperature slope), check this option. Always return to the starting temperature. [X] Return to start temp [?] Reverse scan

All others should be set as for the CD spectral scan.

When the run is finished, follow the shut down procedure or collect a post-melt CD spectral scan to assess refolding Calculate the Tm by fitting the data in python or by fitting the data in Spectrum Manager (truncate the data as needed, calculate the first derivative, calculate the peak)

Powering down After all samples are run, record the voltage at the three reference wavelengths (190, 253.7, 600nm) again and record in log book + total time the lamp was on. To power down, turn off photomultiplier first (right, green). Wait 15 seconds then turn off the lamp (left, red button) Power down temperature controllers if used Let N2 purge for 10 min to prevent ozone accumulation as the lamp cools! Turn off flow adapter box then shut off N2 tank valve. Clean the cuvette - wash with water 5X, ethanol 3X, and leave in 2M HNO3 overnight

For publication, transform your raw mdeg ellipticity:

Other good guides (not necessarily for our software version or machine model): https://chemistry.osu.edu/sites/chemistry.osu.edu/files/Operating%20the%20Jasco%20CD_2_0.pdf https://cmi.hms.harvard.edu/files/cmi/files/cmi_jasco_j-815_cd_getting_started_guide.pdf https://www.cif.iastate.edu/sites/default/files/uploads/Other_Inst/CD/Jasco%20manual.pdf

Expression and solubility test

https://docs.google.com/document/d/15R-14rrS25R5wGI8xIB5GGtr7j11Io-RCvqFQIJrRjo/edit?usp=sharing (please edit as needed)

Expression and Solubility Protocol Goal of this experiment is to test whether or not a designed protein can 1. Be expressed and 2. Soluble in solution. This does not guarantee that a designed protein is functional and expression conditions are optimal. Further purification and characterization need to be done.

Note: this experiment will express protein at 30 C. Proteins may prefer to be expressed at 37 C or 18C. If a protein or protein family member has been expressed before at those specified temperatures, then you can use those temperatures for the experiment, else the optimal temperature and time for expression can be determined in a future experiment. Just because you don’t see a desired band on the gel does not necessarily mean your design can not be expressed.

Materials: Plate that has colonies of BL21 cells with protein of interest LB media (can be found in gel room) (1000x stock) Appropriate antibiotics (can be found in big -20) Culture tubes (can be found in the gel room) 1.5 ml eppendorf tubes Could also use PCR tubes 4x Leammli buffer with BME in it (Laemmli buffer is scattered around the lab) We also have a 2x stock of Laemmli buffer Remember you have to add BME to your own stock. Following the directions on the Leammli bottle to make your own stock IPTG (isopropyl β-D-1-thiogalactopyranoside) (1000x stock) B-per (found on Amy/Eleanor/Lu’s bench) 4-20% gradient Precast SDS-PAGE gels (found in gel room) Things to make the electrophoresis box to run gel (pieces should either be above the sink in the gel room or in a drawer in the gel room labeled as Protein gel Box) 1x SDS Running Buffer (gel room on a shelf shelf next to all the other buffers Solutions for washing, staining, and destaining gels (all in the gel room. Coomassie is next to the microwave. Gel destaining solution is on a shelf text to all the other buffers in the gel room)

Procedure Day 1 Set up a 5 ml LB overnight culture for every protein design that will be tested Pick a single colony Use appropriate antibiotics Let the culture grow overnight in the shaker and to confluence at 37 C at an rpm around 250 Day 2 Remove culter tubes from the shaker and inspect that the cells did grow up If cells did not grow, re-do Day 1 and make sure that the proper antibiotics are used. If it continues to not grow, troubleshooting steps need to be taken and the plasmid needs to be checked (consider colony pcr) Remove 100 ul from each culture tube and transfer into a 1.5 mL eppendorf tube. This will be your uninduced sample. LABEL THIS TUBE. I recommend date, design name, and then U.I. Pellet the cells at top speed either at any table top centrifuge for at least 30 sec. Remove the liquid will trying not to disturb the cell pellet Store this sample in the -20 or -80 until the next day Depending on how the experiment will be done (i.e. which step 5 you do), this sample will either be your uninduced sample on the gel or a backup uninduced sample (Optional) take some sample out and make a glycerol stock Remove 500 ul of cell culture into a cryo tube and add 500 ul of sterile 50% glycerol solution Freeze stock in -80 Spin down the cell culture at 3000 g for 5 minutes and remove the liquid while trying not to disturb the cell pellet Resuspend the cell culture pellets with fresh 5m LB media and make sure the appropriate antibiotics are added back in. Depending on how many designs you are testing, there are two ways you can precede. Use your best judgment on which method you want to use but just know the advantages and disadvantages of each.

5(a). (Proper way) For each resuspended culture tube, split the reaction into two culture tubes. Label one tube as the uninduced sample and the other tube as the induced sample. For the induced sample, add IPTG so the final concentration is 1x.

Example. You do step 4. You label the culture tube as uninduced. You take 2.5 ml of cell culture and place it in a new culture tube and you label that tube as induced. You add 2.5 ul of IPTG to the induced tube. You now have an uninduced tube that has 2.5 mls of cell culture with no IPTG and an induced tube that has 2.5 mls of cell culture that has IPTG Pro: the uninduced sample will be a proper uninduced sample and will have the same OD as the induced sample Con: This will require double the amount of tubes and will require special attention to which tube is which. Example: If you have 10 designs to test this will require 20 properly labeled tubes.

5(b). (Shortcut way) For each culture tube, add IPTG to a final concentration of 1x

Example: You do step 4. You add 5 ul of IPTG to each culture tube Pro: less tubes to deal with. Also less chance of mixing up tubes Con: Technically, the uninduced sample you collected in step 2 won’t have the same OD as the induced sample at the end

  1. Place culture tubes in the shaker at 30 C at 250 rpm and express for 8-16 hours (overnight is fine)

Day 3 Remove culture tubes from the shaker. For each design grab 2 eppendorf tubes and label one tube as induced and the other tube as lysis Example: date GFP I, date GFP lysis For each eppendorf tube, add 100 ul of cell culture to the appropriate tube. (Optional) Take your cell cultures, spin them down, remove the liquid, and store cell pellets in the -20 or -80. You can purify this pellet later to do a mini purification test Spin down your eppendorf tube on a table top centrifuge. Max speed for about 30 secs. Enough to pellet the cells. Remove the liquid. For the lysis samples try your best to remove all the liquid without disturbing the cell pellet Perform a quick cell lysis on the lysis samples using B-per. B-pecr protocol can be found here Resuspend cell pellet with 25ul of B-per solution to each sample Incubate at room temperature for 10-15 minutes After incubation, spin samples at ~17,000 g for 10 minutes Collect the supernatant and place into a new well labeled eppendorf tube Add Laemmli with BME sample buffer (the SDS dye buffer) to your uninduced, induced, and lysis samples For the uninduced and induced samples, have the final concentration of the Laemmli buffer be 2X and the final volume be 100 ul Reason we want a 2x buffer is because we are going to use the high concentration of SDS in the Laemmli buffer to help us lysis the cells for the induced and uninduced. In theory you could just use a 1x buffer but better to be safe than sorry. Example: You have a x4 stock of Laemmli buffer. Resuspend the cell pellet with 50 ul of water. Add 50 ul of 4x Laemllie buffer for a final concentration of 2x with a final volume of 100 ul. Vortex solution SEC suggestion: Resuspend the pellet in 16 uL water and 16 uL Laemmli buffer so that the volume/concentration is matched to the ‘lysis’ sample. If you load 10 uL of all samples on the gel (~30% of sample) you should be able to semi-quantitatively compare levels of your protein between samples. For the lysis buffer, have the final concentration of the Laemmli buffer be 1x and the final volume be ~32 ul. Example: You have a 4x stock of Laemmli buffer and the volume of the lysis sample is ~25 ul. Add 8 ul of Laemmli buffer to the lysis solution. Vortex solution. Place samples on a 95-98C heating block and let samples denature for about 15 minutes Thermo says you can denature for 5 minutes but since we want to lysis the cells I have found that 15 minutes works During this time set up gel box for electrophoresis Grab a Pre-casted 4-20% SDS-PAGE gel and assemble the electrophoresis box Here is a step by step guide to set up the box. Go to section 2.2 on page 9 Key points are: Wash the precast gel, REMOVE GREEN TAPE AT BOTTOM OF GEL, remove the combs, wash the wells, and make sure that there are no leaks, and fill After samples have denatured, do a quick vortex and quick spin to collect any evaporated solution in the tube When samples are ready and gel box is set up, load Protein ladder and samples on the gel 5 ul of Precision Plus Protein Dual Standards (green top, and it is kept in the small fridge in gel room) 10 ul of your sample Suggested layout: Lane 1: Ladder, Lane 2: Design 1 uninduced, Lane 3: Design 1 induced, Lane 4: Design 1 lysate, Lane 4: Design 2 uninduced… Run the gel at 125V for 1 hours or until the dye front reaches the bottom of the gel Bio-Rad says you can bump up the gels to 300V and run it for 20 minutes but please be careful if you are going to do this. THis may cause the gels to heat up and melt and may cause smearing of bands Wash, Stain, and Destain gel Here is how to break open the gel cassette (go to step 5) Here is a basic protocol of washing, staining, and destaining gel Ask someone in the lab how they do things. There are more than one way to stain and destain gels. Choose your own adventure FQA How do I read this gel/how do I know things worked? First you have to know the molecular weight of your protein design (remember that you will probably have an affinity tag and cut site and this adds weight). If you see a thick band in the induced lane that is about the right size as your protein and you don’t see a thick band in the uninduced lane, then this is evidence that your designed protein can be expressed. If you see the same thick band at the same size in the lysis lane then this is evidence that your protein is soluble. THIS IS GREAT! I see a band in the uninduced lane that is about the right size for my protein. Is this my protein? !Yes! This is called leaky expression and is common with BL21 DE3. As long as you see the same band in the induce and lysis sample, and those bands are thicker, you are all good I am struggling to find a band that is the right size for my protein of interest/I think I see a band but it is not thick. Is my protein there? Maybe. If you are struggling to determine if your protein is expressed then it could be three things. 1. Your protein can not be expressed 2. Your protein’s expression level is really low 3. You forgot to add IPTG. This experiment is not definitive if a protein can be expressed or not. To troubleshoot I would do a mini protein purification with the sample you saved in step 4 on day 3. You may find out that your protein did express just at low levels and you may need to optimize your expression. Consider expressing at different temperatures. I see a band that is the right size for my protein in the induce sample but I don’t see the band in the lysis sample. Where did my protein go? This result could be two things. 1. The protein aggregated and is in inclusion bodies 2. Something went wrong with the lysis step. I would take your sample from step 4 day 3 and redo the lysis step. Either consider adding more B-per or use sonication. After you lysis, spin, and collect the supernatant of your sample, I would run a gel where one lane is the lysis pellet and the other lane is the lysis supernatant. If you only see your band in the lysis pellet lane then your protein has aggregated. Troubleshoot accordingly

TL:DR Procedure Day 1 Start overnights in LB with appropriate antibiotics for all designs in culture tubes. Make sure designs are in BL21 DE cells Place culture tube at 37 C 250 rpm

Day 2 Remove culture tubes from shaker and collect 100 ul of cell culture from each design and label tube as uninduced Pellet cells, remove solution, and keep in -20 Pellet culture tubes at 3000 xg for 5 min Remove solution, and add 5 ml of fresh LB and appropriate antibiotics Add 5ul of 1000x IPTG to culture tubes Place tubes in shaker set at 30 C 250 rpm and express for 8-16 hours

Day 3 Remove tubes from shaker For each design collect two 100 ul cell culture samples. One tube will be your induce sample and other tube will be your lysis sample Pellet both samples and remove liquid Lyse the lysis samples Resuspend cell pellet with 25 ul of B-per Incubate at Room temp for 10-15 min Spin at ~17,000 g for 15 minutes Collect supernatant Denature uninduced, induced, and lysis samples at 98C for 15 minutes Uninduced and induced samples should have a final volume of 100 ul and 2x Leammli with BME buffer Lysis sample should have a final volume of 32 ul and a 1x Laemmli with BME buffer Load samples on the gel 5 ul of Protein Ladder 10 ul of protein samples Load samples on the gel and run at 125 V for about an hour or until dye front has reached the bottom of the gel Wash, Stain, and Destain gel

Chemical transformation and glycerol stocks

https://docs.google.com/document/d/1yvBgWy5qZKOD9QvX9V2dHYkPLgI8CpZb00zXySUkbJk/edit?usp=sharing (please edit as needed) Chemical transformation and Glycerol Stocks/Overnights

This protocol is based off of this website (this all around a great website and should be looked at to learn more about the whole process of bacterial transformation): https://www.thermofisher.com/us/en/home/life-science/cloning/cloning-learning-center/invitrogen -school-of-molecular-biology/molecular-cloning/transformation/bacterial-transformation-workflow.html#:~:text=With%20chemical%20transformation%2Cchemically%20competent,minutes%20in%20a%20polypropylene%20tube.

Note: that most timing steps in the protocol can be flexible except for heat shocking!

Material: Ice Chemically comp cells (can be made or purchased) DNA (concentration 100 ng/ul or higher) S.O.C media or LB media Heat shock bath set @ 42C Culture tubes Appropriate antibiotics (for overnight cultures) LB plate with appropriate antibiotics Cryo tubes (for glycerol stock) 50% sterile glycerol solution (for glycerol stock)

Chemical Transformation Protocol Grab chemically competent cells from the -80 freezer and thaw on ice If doing a DNA mini prep later or making a glycerol stock grab Top10 cells If expressing protein grab BL21(DE3) cells You really only need about 25 ul of competent cells to do a transformation If grabbing cells from the switch comp cells, each tube should be about 55 ul which is enough for two transformation so you can split the cells if doing more than 1 transformation Add 1 ul of DNA to thawed cells and let it equilibrate for ~ 30 mins on ice If DNA concentration is lower than 100 ng/ul considered higher volume of DNA Do step 4 while equilibrating Grab plates with appropriate antibiotics and warm them in the 37 incubator Place cells with DNA in the heat bath set at 42C and let it sit for 45 sec. Recover cells on ice for 2 min Add 1 ml of S.O.C media to cells and tape it in the shaker set at 37 C at rpm 250 and let it recover between 30 min to 1 hour If you don’t have S.O.C media use LB media Grab plates from incubator and place sterile glass beads on the plate Min: 5 beads Max: 10-15 beads Pipet up and down recovered media and plate 120 ul of solution Place plate in the incubator @ 37 for overnight growth Pull plate out the next day Do not let plate grow longer than 24 hours

Making Glycerol Stocks This procedure is based of of this link

After a transformation, it is always a good idea to make a glycerol stock. This can be used as a back up and also a place where you can go to make more plasmid (so you don’t have to transform again in the future). In this lab, we make glycerol stocks of top10 cells with your plasmid of interest. Reason being is that when you leave this lab and someone needs to re-make your plasmid, they can go to this glycerol stock, make more plasmid and then proceed with whatever. All this to say MAKE A GLYCEROL STOCK

Procedure: Day 1 From the plate you transformed above, pick 1 colony and start a 5 ml overnight culture in LB media with the appropriate antibiotics You can take a sterile pipette tip, scrap one colony and drop it in a culture tube Set in the shaker at 37 C 200-250 rpm and let it grow overnight. DON'T LET IT GROW FOR MORE THAN 24 HOURS Day 2 Grab the appropriate amount of cryo tubes and CLEARLY LABEL YOUR TUBE I suggest, Initials, Date, name of design, plasmid vector, and cell line the plasmid is in Ex. DWG, 2/2/22, LUCS 32, pet28a, Top10 The plasmid vector from TWIST tells people what antibiotic to use and an idea about the affinity tag, its location and if it has a cut site. Cell line tells people if you should express protein or express plasmid Top10 = plasmid BL21 = protein Using sterile technique (i.e. next to a flame), add 500 ul of 50% of sterile glycerol to each cryo tube People have used glycerol stocks ranging from 50%-25%. The key is that you want your final glycerol concentration to be around 15%-25%. So make the stock whatever you want. Just know that 100% glycerol is really viscus Still using sterile technique (i.e. next to a flame), add 500 uL of your culture sample into the appropriate cryo tube making sure to pipette up and down Store well labeled cryo tubes in the -80. With the reset of the culture tube you can mini prep (also just google qiagen miniprep protocol and the first link is a pdf of the mini prep protocol), and then send the plasmids out for sequencing to check

Affinity protein purification

https://docs.google.com/document/d/1_HHqOZmUGwI5xWMWUvjgHrf9rqVGrqivmdkO2dXWsFI/edit?usp=sharing (please edit as needed)

TABLE OF CONTENTS

Affinity Protein Purification through immobilized metal affinity column (IMAC) 1 Background: 1 Lysing your sample and getting a Supernatant 2 Materials 2 General Procedure 3 General Tips people in the lab have found 5 Amy’s protocol for large batch His-tag purification: 5 Dominic’s protocol for purifying 5 mL culture tubes 6 Catherine’s protocol for Strep-tag purification 8

Affinity Protein Purification through immobilized metal affinity column (IMAC) The principle and general steps in this protocol can be applied to any affinity based purification. Your buffers will be different though.

Background: This protocol is going through IMAC purification. In our lab this is typically done using Ni-NTA resin and assumes that your protein has a 6x-HIS tag. It is really important that your protein has a 6x-His tag because histidine will bind to nickel, so long as you are above Histidine’s pKa. No neutral histidine, no binding.

There are four basic steps to any affinity-based purification. Prepping and equilibrating the resin/column Loading the resin/Column with supernatant Washing the column to remove any unwanted protein and any nonspecific binding to the column Eluting your protein off the column

For IMAC, the way we wash and elute our resin/column is to use different concentrations of imidazole (again at a certain pH). Imidazole looks like histidine, and therefore will compete to bind to nickel. A higher concentration of imidazole means it will out compete the His-tag and therefore your protein of interest will “elute” off the resin/column.

Why are you telling me all of this? IMAC purification is a very useful protocol, but sometimes it does not work. The usual culprits of why it does not work are:

  1. Your concentration of imidazole is off: a. Too high and your protein is coming off too early. b. Too low and your protein will stay on the resin.
  2. Your buffers are not pH properly. a. Both imidazole and histidine need to be above their pKa in order to bind to nickel
  3. Mixed up the buffers, and eluted too early (this pretty much follows 1.) a. If this is the case you can still recover! Just dilute your samples with wash buffer and throw it back on the column.

Lysing your sample and getting a Supernatant This protocol is not going to go over how to lysis your protein. The reason being is that lysing is very dependent on the protein you have and how big your culture volume is. It is up to you to know how to lyse your protein. In our lab there are three main ways of lysing. 1. Chemical detergents 2. Sonication 3. Microfluidizer. However, there are many other ways to lysis your protein. Research on which protocol you should use. If using the sonicator or microfluidizer, please consult whoever is in charge of it to learn how to use these machines if it is your first time using it.

Once you have lysed your samples, you have to spin it down. ~18,000 x g for 30-45 min at 4 C should do the trick. Your supernatant must be clear before moving on! If it is not clear then transfer the liquid to a new tube and spin down again. You can either use the table top centrifuge to spin down samples. This takes the 1.5-2 ml tubes. We also have a floor centrifuge that is shared with the Krogan lab. The floor centrifuge uses 50 mL tubes. Please ask whose lab responsibility it is that oversees the floor centrifuge to see how to properly use it.

Materials Ni-NTA resin This is kept in the refrigerator in the gel room Remember that resin is kept in ETHANOL!! And that it is half the volume This means if you want 1 ml of resin you have to take up 2 ml slurry (resin + ethanol) Also means you have to wash the resin with water first! Gravity column or falcon tubes (or 1.5 ml tube) Gravity column = gravity column purification (explained below) Falcon tubes = Batch purification (explained below) Volumes of columns and tubes is dependent on how much resin you have Equilibration Buffer Low to none Imidazole + salt + Buffer Example recipe: 50 mM Tris-HCl pH 7.5, 300 mM NaCl, 10 mM Imidazole pH 8 Your buffer can be different depending on what you want Wash Buffer Low Imidazole + salt + buffer (but higher than equilibration buffer) Example recipe: 50 mM Tris-HCl pH 7.5, 300 mM NaCl, 25 mM Imidazole pH 8 Your buffer can be different depending on what you want Elution Buffer High Imidazole concentration + salt + buffer Example recipe: 50 mM Tris-HCl pH 7.5, 300 mM NaCl, 300 mM Imidazole pH 8 Your buffer can be different depending on what you want Tubes to collect each step Ice When your sample is not on the resin you should keep it on ice as much as possible Cold prevents aggregation

General Procedure This assumes you have lysed your samples, spun down the lysis, and have clear supernatant. There are two ways to purify. One is to use a gravity column. The second is to do batch purification.CLICK ON THIS PROCEDURE by Thermo to look at the difference. Both ways have pros and cons and everyone has a preference. Read through the thermo protocol and talk with people in the lab to see which way will suit your needs. Regardless of which method you use, both gravity column and batch basically follow the same steps.

Another important note is that most purification protocols are based on bed volumes. A bed volume is how much volume of resin you have. For example, if I have 250 ul of resin that will equal 1 bed volume. When the protocol says to wash resin with 4 bed volumes that is equal to 1 ml (250 ul x 4 = 1 ml). The reason we do this is because the volume of the buffer you use is dependent on how much resin you use. How much resin you use is very dependent on how concentrated your sample is and how much sample you have. Ni-resin has a binding capacity of 60 mg/ml (this is a lot). There is no general rule on how much resin you need but here are some suggestions. THIS IS NOT PERFECT AND IS VERY VERY DEPENDENT ON YOUR SAMPLES!!!! Use your best judgment and past experiences.

5 ml culture volumes = ~125 ul resin 50 ml – 100 ml culture volumes = ~250 ul resin 100-500 ml culture volumes = ~ 500ul - 750 ml resin 1 L culture volumes = ~1 ml resin

RESIN CAN BE REUSED! Before throwing away resin, remember that you can reuse it! You can either wash it a lot with an elution buffer, then wash with water, and store it with 20% ethanol and keep it cold. This is really useful if you know you are going to do future purification of the same protein. You can also strip and recharge resin. Look at the thermo protocol to learn how to do this. Don’t just throw away resin!

(do steps 1 and 2 while you are lysising and spinning your samples) Wash resin with 2 bed volumes of water to remove ethanol Discard liquid Equilibrate resin with Equilibration Buffer for 2-3 bed volumes Discard liquid Add supernatant to resin If you have a lot of sample add it to the resin in batches Collect liquid if you would like to put on a gel Add 5-6 bed volumes of equillibration buffer to the resin Do this in batches. Don’t add all the liquid at once Collect liquid if you would like to put on a gel This step is optional but highly recommended. The main idea here is to essentially wash away all protein that will not bind to the column (i.e. equlibrate the column) leaving only specific and non-specific protein on the column. This just helps give you cleaner product in the end but you do run the risk of opentially losing some product. Use your best judgment. Add 5-6 bed volumes of wash buffer to the resin Do this in batches. Don’t add all the liquid at once Collect liquid if you would like to put on a gel Main idea is to remove non-specific binding Add 3-5 bed volumes of elution buffer to resin COLLECT LIQUID!!! This should have your protein of interest Do this in batches, and keep samples separated at first until you have run a gel This will save you time later when you concentrate. For example, you did 5 bed volumes and collected each bed volume in a separate tube. The gel shows that only tubes 1-3 have protein. You can toss tubes 4 and 5 and that is less volume you have to concentrate Run a gel of collected samples to see if you have protein, how clean it is, and qualitatively how concentrated it is.

General Tips people in the lab have found If doing batch protocol, let the supernatant bind to the beads before spinning. Some people have let supernatant incubate overnight in the cold room Some people also do this for gravity columns If doing the batch protocol, remember to resuspend the resin once you have removed the supernatant and are adding a new buffer. If measuring the 280 signal, please be careful interpreting the 280 signal. Imidazole can absorb around 280 nm and since your elution buffer has a high concentration of imidazole this can throw off your 280 reading. Long story short if you see 0 absorbance THIS DOES NOT MEAN YOU HAVE NO PROTEIN. It might just mean your protein concentration is lower than the imidazole concentration and is getting washed out. ALWAYS RUN A GEL.

Amy’s protocol for large batch His-tag purification:

Materials Equilibration buffer (50mM Tris pH=7.5, 300mM NaCl, 10 mM imidazole) Wash buffer (50mM Tris pH=7.5, 300mM NaCl, 25mM imidazole) Elution buffer (50mM Tris pH=7.5, 300mM NaCl, 250mM imidazole) - not needed if doing enzymatic cleavage Enzyme cleavage buffer - depends on your enzyme Ni-NTA resin Enzyme capture kit (ideal, but not needed if sample is only needed for the next few days) Protein storage buffer - depends on downstream applications

Protocol Spin down cell pellet (8000g, 10min) and optionally freeze at -80C Lyse the cells with sonication or detergent and add a protease inhibitor cocktail pill. Amy strongly prefers B-PER/Y-PER detergent-based lysis - sonication can result in a lot of denatured protein and higher MW impurities. Spin down lysate 18000rpm, 10min While lysate is spinning down, prep your Ni-NTA resin. Add 2mL Ni-NTA resin slurry per 1L culture to a centrifuge-compatible tube Add at least 2 resin bed volumes (aka half of the slurry volume) milliQ water and spin down 750 rpm 2-3min + decant to remove EtOH (decant either by carefully pouring out supernatant or pipetting) Add at least 2 resin bed volumes of equilibration buffer, resuspend, and centrifuge + decant Mix lysate supernatant with an equal volume of equilibration buffer (it’s ok to get some of the cloudy liquid - no amount of centrifuging will settle it) Pour supernatant onto beads and let proteins bind - 30 min-1 hour on a nutator at 4C ideally Wash pellet 3X in 2 resin bed volumes of wash buffer (centrifuge 750g, 2-3min)

For enzymatic cleavage: Wash pellet 1X in 2 resin bed volumes of enzymatic cleavage buffer (centrifuge 750g, 2-3min) Follow instructions from enzyme kit to cleave the His-tag off of your protein. Amy recommends doing a small scale test digest with 10ug of protein immobilized on the Ni-NTA beads in each condition (4x, 2x, 1x, 0.5x units of enzyme for 2h, 4h, 8h, 20h) because every protein is different. I have had to use up to 4x for 20h to get efficient cleavage. Prepare a fresh mL of Ni-NTA resin (wash with water, protein storage buffer) Centrifuge cleaved protein solution and fresh resin separately Repeat 3X: Add supernatant from the cleavage reaction onto fresh Ni-NTA resin and invert to mix. Centrifuge and transfer to a fresh tube. Add 1 resin bed volume of protein storage buffer onto cleavage resin to increase yield Centrifuge “new” resin 2500 rpm, 5min and transfer the supernatant to a fresh tube If using a cleavage capture kit, mix with cleavage enzyme immobilizing resin now.

For eluting with the tag intact (not recommended unless you know for certain the tag does not interfere with protein structure/function): Elute 3X with 1 resin bed volume of elution buffer.

Filter all of the collected supernatant from the fresh tube with a syringe into another clean tube Concentrate/buffer exchange into your protein storage buffer with an appropriate MWCO Amicon column. Depending on how many spin downs you do, this may take a couple hours.

Dominic’s protocol for purifying 5 mL culture tubes Lysising samples: This will depend on how many samples you have. I would recommend making the lysis buffer right before you have to lysis and to make extra. Below is an example of making a 20 ml lysis buffer for 18 samples. Adjust the volumes as needed.

20 ml Lysis buffer (1 ml per sample 18 samples) - (OPTIONAL) .2 ml of Lysozyme (100 mg/ml stock) final concentration 1 mg/ml. Please be careful when using lysozyme. Lysozyme has a MW of 15 kDa. So if your protein is around that size and you don’t wash the column that well then you can end up having lysozyme sticking around. Please use your best judgment. - .2 ml of MgCl (1 M) final concentration 100 mM. This is needed for DNase - 2 ul of DNase (Could also use Benzenase which is a really really effective enzyme you only need a small amount of it. Please do not use a lot of it. 1U is defined as 37 ug DNA per min that is a lot. Use your best judgment.) - 19.6 ml of B-Per in Phosphate - total Volume ~ 20 ml Resuspended spun down culture tubes with 1 ml of lysis buffer and placed in a new eppendorf tube. Incubated samples at 30 C for ~20 minutes occasionally agitate samples Spin samples at max speed on table top for 30 minutes at 4 C Collecture supernatant (super) (optional but recommended) take 15 ul of super to be run on a gel Move on to purifying

Purifying sample:- Suggested MNC edits:

  • Take pellet out of freezer, add protease inhibitor tablet, add ~500 uL (?) B-PER for 5 mL culture
  • Resuspend and dissolve tablet/pellet by pipetting
  • Transfer to 1.5 mL eppendorf
  • Incubate 10-15 minutes RT
  • Spin 10 minutes max speed
  • prepared batch tubes with 150 ul of Ni-NTA resin (bed volume), washed with water and equilibrated with Equilibration buffer
    • Put resin in mct (microcentrifuge tube), add 2 bv water, spin 750 g 2-3 minutes, aspirate, add 2 bv equilibration buffer, spin 750g 2-3, aspirate
  • Loaded lysate on resin, let them incubate on resin in cold room on a shaker for 15 minutes
  • spin samples down at 800 g for 3 min (ANY TIME I SAY SPIN DOWN FROM NOW ON THIS IS THE PARAMS), removed super
  • 1 ml of Equilibration Buffer to each tube and shake back and forth (6.6 bed volumes washed)
  • spin samples and remove super
  • 1 ml of Equilibration Buffer to each tube and shake back and forth (13.3 bed volumes washed)
  • spin samples and remove super
  • 1 ml of Equilibration Buffer to each tube and shake back and forth (20 bed volumes washed)
  • spin samples and remove super
  • 1 ml of Wash buffer to each tube and shake back and forth (6.6 bed volumes, 26.6 washed columns total)
  • spin samples and remove super
  • 1 ml of Wash buffer to each tube and shake back and forth (13.3 bed volumes, 33.2 washed columns total)
  • spin samples and remove super
  • 1 ml of Wash buffer to each tube and shake back and forth (20 bed volumes, 39.8 washed columns total)
  • spin samples and remove super
  • 500 ul of Elution buffer and shake back and forth (3.3 bed volumes)
  • spin samples and collect super
  • 500 ul of Elution buffer and shake back and forth (6.6 bed volumes)
  • spin samples and collect super
  • 500 ul of Elution buffer and shake back and forth (9.9 bed volumes)
  • spin samples and collect super
  • 500 ul of Elution buffer and shake back and forth (13.3 bed volumes)
  • spin samples and collect super

Catherine’s protocol for Strep-tag purification The general protocol from the strep-tactin resin manufacturer can be downloaded here: https://www.qiagen.com/us/resources/resourcedetail?id=0fadc040-86ad-4e31-bb58-7e83ceb77b72&lang=en I generally follow the “Protocol: Batch Purification of Strep-tagged Proteins Using Strep-Tactin Superflow Plus”

Materials Needed: Binding Buffer Elution Buffer Regeneration Buffer Strep-tactin Resin Gravity flow column

Buffers needed (store these in the cold room): Binding Buffer: 100mM Tris-HCl pH 8, 150mM NaCl, 1mM EDTA To make 1 L of buffer: 30mL of NaCl of 5M NaCl 100mL 1M Tris-HCl, pH 8 2mL of 0.5M EDTA 868mL water Elution Buffer: 2.5mM desthiobiotin (DTB) in binding buffer (0.536mg DTB per 1mL of wash buffer) this can be kept at room temp for ~ 1 day so should be made fresh for each purification Regeneration Buffer: 1mM HABA (2-[4’-hydroxy-benzeneazo] benzoic acid) in binding buffer

Purification: Prepare buffers. I usually make the binding buffer ahead of time (and it can be stored for a while at 4C). 1 L should be sufficient. To prepare elution buffer, add 0.536mg of desthiobiotin per 1 mL of buffer. This should be made fresh on the day of purification. Lyse cells following whatever lysis method works for your cells (which can depend on things such as culture volume). While spinning lysed cells, wash strep resin (if using new resin) Add about ~125 uL resin for a 5mL expression or ~2mL of resin for larger scale expressions to a centrifuge-compatible tube. Wash resin with 2 CV (1 CV = amount of resin you are using) of water to remove ethanol and spin down 750 rpm 2-3min. Discard liquid. Equilibrate resin with 2 CVs binding buffer, spin down, and discard liquid. Incubate the lysed sample with washed resin. Ensure cellular debris in your lysed solution is FULLY spun out before adding it to the resin. The lysis should not be cloudy at all as this will make purification MUCH slower. collect 15uL of liquid for a gel. Add lysis to a tube with resin. Let proteins bind ~1 hour on a nutator at 4C. After incubating, load the resin + lysis solution into a column with the bottom capped. I do all my purification in the cold room as the resin manufacturer’s protocol suggests doing everything at 4C, though it may not be necessary. Remove the cap and collect flow-through. Wash column with ~4x4 CVs binding buffer (to remove non-specific binding) collect each wash fraction for gel (this will tell you if you are doing a sufficient number of wash steps to remove non-specific binding to the column/if you are losing a lot of your protein). You can do more wash steps for cleaner protein. Add 6x0.5 CVs elution buffer and collect fractions for each elution round. Collect each elution fraction. You can regenerate the strep column resin up to 5 times. To do so: Make regeneration buffer: 1mM HABA (2-[4’-hydroxy-benzeneazo] benzoic acid) in binding buffer, which is ~24.2mg HABA per 100mL binding buffer. Make pH 10.5 wash buffer (this helps remove the HABA). Run 3x5CV regeneration buffer through column. The resin should go from an orange color when you first add the regeneration buffer to a deep red color (this means the HABA has replaced the DTB). Run 2x5CV wash buffer. Run 2x5CV wash buffer at pH 10.5. This step should really help remove the color from the resin (the goal is to get the resin back to the initial white color. If it’s still pink, run a few more CV’s through it until it clears up. I also find pipetting in the column up and down a few times after adding the wash buffer helps). Do not leave the resin in the pH 10.5 buffer for too long as it should not be left at a high pH. Run 2x5CV wash buffer. At this point, the resin should be white again. Store resin in 2CV wash buffer at 4C. (It can also be stored in the regeneration buffer if you don’t have the time to do all of the regeneration steps).

Ordering sequencing in Elim

https://docs.google.com/document/d/13P1Ic7cPaN3fHtChqDd_7kh_75eVxQmwQ_TVwXucj2g/edit?usp=sharing Ordering Sequencing in Elim

General answers to questions can be found on this website: https://www.elimbio.com/services/dna-sequencing/

Go to https://www.elimbio.com/ If you do not have an account go to this link and fill out the form https://order.elimbio.com/onlineorder/registration.asp Click on Sequencing -> Fill out a New/Pending Order Form (where the red circle is)

Then click on “To create a new order form with a new order number. Please click here” Fill out the two questions as needed WHen you click “Online ORder Form” It will ask you have many samples and if they are Premix or Non-Premix. If you are going to use Elims primers click on the “non-premix” tab For the PO there is a table in the lab of which great you should be pulling from and there is a special PO number for Elim (and quintara). Use the appropriate PO. If you do not have this table or have no idea what this is, ask someone in the lab and they can forward you the email Fill out the form as needed If using Elim’s primers click on the “Elim common primer” a new window should pop up and click on the primer you want to use. Go to this website to look at the primers that Elim. If you are going to use a specific primer a lot in the future you can contact Elim and they can have your primer in stock. We have some on file. If this interests you, reach out to Elim. There are these Green down arrows that will autofill a column to whatever the first row is. This is nice to have when you have a lot of samples Once you are done filling out the form, click on submit, then there is an order verification page. If you are still happy with everything then click “Submit ORder Form now” You will be given a confirmation page. YOU NEED TO PRINT OFF THIS PAGE!! I would print two copies. One copy for your records, other copy to be placed with your sample Sample prep!!: Place DNA samples in a PCR tube Elim would like 50 ng/ul of 10 ul of DNA sample for plasmids. I give them 12- 15 ul just in case. Number the PCR occurring to how you filled out the form. Ex. The 1st sample on the form is the first PCR tube so that PCR tube should be numbered 1 etc. Place samples and confirmation page from step 9 in a clear bag. Make sure the barcode is showing. Place the bag in the Elim folder outside of the wet label in 309. YOU'RE DONE! YAY! Results should come in within 24 hours if you place your samples in before 3 pm (I think)

Ordering from Twist

https://docs.google.com/document/d/1HGlJmbuJ454yYQbzGJEwpcF238NgIqk46xAlqrZPxJM/edit How to order protein designs from Twist for bacterial expression Input: the amino acid sequence of your design (and potentially your vector if you are not using one of Twist’s vectors)

Make sure before ordering designs you have the following: A folder with the PDBs of all your designs A document describing in detail your design protocol and selection rationale A spreadsheet with all of your design amino acid sequences (Generated by Twist - not needed beforehand but keep this with your other records) A folder of all your plasmid maps Upload all these documents to Box!

Create a Twist account and email our Twist representative ([email protected]) to be added to the Kortemme lab group Sign into your Twist account Click “Genes” (if you are ordering designs on a small scale, i.e. not a library)

Name your project and click “Start New Project”

If you are ordering designs pre-cloned into an expression vector, select “clonal genes”. If you are ordering gene fragments to do the cloning yourself, select “gene fragments”.

Select “amino acid sequence” and your import method of choice

Choose E coli for the codon optimization table in the dropdown menu

Put an asterisk at the end of your sequence to signify a stop codon - they will NOT put one automatically

To change the vector or add flanking bases (if your design is too short) for all designs, select the box for all designs and click the Change Vector or + Flank For the flank sequence, I don’t think it matters too much what this is as long as there are not a lot of homologous repeats/secondary structure. Sometimes I will just take a portion of the non-coding plasmid backbone of a different vector. Add a cut site at the beginning of the flank (so you can still digest out your design if you need to) and put the flank after the stop codon (at the 3’ end). For the vector, many of us have been using pET28a+ with an NdeI/XhoI cut site. It has the following structure: MET – 6X His tag – thrombin cleavage site – NdeI – [your design] – XhoI cut site – 6X His tag so you do not need to put the cut sites in manually.

Download the genbank files of all your sequences Double/triple check these in a plasmid viewer of your choice (e.g. Benchling) and keep them for lab records.

Click “Continue” once you are sure the translation of your protein is correct (in frame, right sequence, all tags are present, etc.) and the complexity is “standard”. Select the “no additional fee” options. If you are just ordering a couple of designs, you can have them shipped in individual 2mL tubes for free. Otherwise I tend to use the 96-well PCR plate (I find it easier to pipette the samples from these). Typically I select “send everything” since the amount is usually varied regardless.

Click “Request Quote” Check out. Select the appropriate PO number for your project (check with Tanja if you are unsure) and use the following shipping address:

Agree to the terms/conditions and submit your order! DNA usually will be in the lab after 2 weeks.

Spotting yeast from liquid culture on a plate.

  • PCRs of yeast DNA (genomic or plasmid) work best from colonies. If your yeast is already on a plate, skip to (2).
  • adapted from Buratowski Lab Wiki

Steps:

  1. Get some cells from exponentially growing culturesm say 5ml at OD600~0.8
  2. Spin down (5 min, at least 2k rpm in tabletop centrifuge)
  3. Re-suspend in 1ml ddH2O to wash, then re-suspend in 500 ul
  4. Prepare serial dilutions if analyzing growth phenotypes
  5. Spot 10 ul of each sample into a (labelled) spot on a grid.
  6. Dry the plates 10-20 min at least. Can speed up by placing plates lid-off in a laminar flow or beside a lit flame.
  7. Incubate

Yeast Transformation with LiAc

Prepping for Yeast Transformation

  1. Start a 5 ml cell culture by transferring 5 ml of yeast media (e.g. YPD) into glass conical tube(s) and inoculating the conical tube(s) with a single colony
    • Note: If yeast cells are grown in media without antibiotics, use careful sterile technique to not contaminate the cultures.
  2. Grow the yeast cell cultures overnight at 30°C on the ferrous wheel incubator or shaker.
  3. The next morning, measure the O.D. of the cell culture(s) on the Simple Reads program at 600.00 nm and dilute the cell culture(s) down to a starting O.D. of 0.25. Dilute the cell culture(s) with the yeast media that was used to grow the cell cultures the night before.
  4. Grow the diluted cell cultures back up to an O.D. of 0.7 (this usually takes about 4 hours).

The Transformation

  1. Transfer 50 ml of the cell cultures into conical tube(s) and centrifuge the tube(s) for 3 minutes at 3000 rpm. Be sure to use all of the cell cultures!
  2. Pipette the supernatant out of the tube(s) and discard it.
  3. Re-suspend the pellet(s) in the conical tube(s) with 800 µl of H2O.
  4. Transfer the re-suspended pellet(s) into eppendorf tube(s) and centrifuge the tube(s) for 5 minutes at 3000 rpm. 5.Discard the supernatant and estimate the volume of pellet formed in a single tube. Match that pellet volume to the volume of H20 needed to re-suspend the pellet(s).
  5. Pipette 50 µl of the re-suspended cells into new tube(s). This transfer contains 25 µl of cells and 25 µl of water.
  6. Spin the tube(s) down again for 1 minute and 30 seconds at 3000 rcf. When centrifuge is complete, discard the liquid . 8.Then, to each tube, add ingredients:
    • 36 uL of 1M LiAc
    • 10 uL ssDNA carrier (2 mg/ml), prepared by boiling for 5 min then placing on ice
    • 1 uL of plasmid (or 15 uL of PCR product)
    • 59 uL of H2O (or 44 uL if using PCR product)
    • NOTE: LiAc "loosens" the cells' membranes to allow the DNA into the cells. ssDNA aids the transforming DNA into the cells so transformation can take place within the cells
  7. Next, re-suspend the pellet of cells gently
  8. After, pipette 240 µl of 50% PEG 3350 in the tube(s) and VORTEX the tubes. Incubate 30 min at RT.
  9. Add 50 uL DMSO after incubation is complete.
  10. Incubate the tubes in a 42C water bath for 40 minutes to heat shock.
    • Note: PEG is very viscous and thus, if it is added first, it becomes very difficult to re-suspend the cells evenly throughout the mixture, making transformation of the cells very difficult. PEG helps the cells stay in the mixture throughout, while minimizing gravity's affect.
  11. Once the last step is finished, spin the tubes down once more for 3 minutes at 3000 rpm.
  12. Dump the supernatant and re-suspend pellet(s) in 350 µl of yeast media.
  13. Incubate cells at 30C for 4h to rescue from heat shock (this is critical)
  14. Then, plate the cells on desired selective media and grow in a 30C incubator for approximately 3 days (or more).
    • NOTE: Do not need to plate all of the cells, a quarter or a sixth is plenty.
  15. After 3 days have elapsed, proceed to Yeast Colony PCR.

Yeast Colony PCR

  • according to Tina, "the trick for good colony PCR with yeast is fresh yeast that is not overgrown and has never been in the fridge"

Steps:

  • Aliquot 40 ul of 20 mM NaOH into a tube
  • Lightly touch a colony with a pipette tip, re-suspend in the NaOH. Solution should be slightly cloudy but still transparent. If it gets too opaque, you have too many cells and you'll have too much DNA for the PCR
  • Incubate 20 min at 95C
  • Spin down until you can see the cell pellet (can just use a small tabletop spinner, or a microcentrifuge)
  • Use 1 ul of supernatant as a template in a PCR, 35 cycles
  • Take 1 ul of PCR product, use as template for another PCR, 35 cycles
  • Run gel to confirm band size

PCR recipe:

Reagent Volume (uL)
2x Master Mix 10
template 1
10 uM Rev primer 1
10 uM Fwd primer 1
H2O 7
TOTAL 20

PCR Protocol:

  • 98C, 3 m
  • Repeat 35x:
    • 98C, 30s
    • Ta, 30s
    • 72C, 60s
  • 72C, 2m

Introducing insertions/deletions using Q5 master mix

Lu Hong

2021/02/05

  1. Design your primers using the NEBaseChanger. Note the provided annealing temperature Ta in particular.

    • For deletion, the program will design primers that linearize your plasmid, excluding the region to be deleted; for insertion, the insert will be attached to the 5' end of the forward primer (for ≤ 6 nt inserts) or split between the 5' ends of both primers (for > 6 nt inserts). The insertion length is limited by primer synthesis.
    • The basic idea is to generate a correct linear sequence using PCR, and then blunt-end ligate with a kinase and ligase.
    • More explanation is provided in the Q5 Site-Directed Mutagenesis Kit product information page.

  2. Set up the reaction mixture (see also this page and this page); bring the total volume to 25 μL with nuclease-free water (ddH2O is sufficient in practice).

    • For a 4-5 kb plasmid, I usually use 10 ng of template. It may be necessary to setup a concentration gradient to optimize the condition for new plasmids.
Component Target
Q5 2x master mix (NEB catalog # M0494S) 1x
Forward primer 0.5 μM
Reverse primer 0.5 μM
Template DNA 1-25 ng
  1. Set up the PCR reaction
Seg. # Cycles Temp (°C) Duration Note
1 1 98 30 sec Denature template
2 25 98 10 sec
2 - Ta 10-30 sec
2 - 72 variable 20-30 sec/kbp (< 6 kb) or 40-50 sec/kb (> 6 kb)
3 1 72 2 min Final extension
4 1 4-10 Hold
  1. Set up the KLD (kinase-ligase-DpnI) reaction; incubate at 37 °C for 30 min.
    • NEB also sells premade KLD enzyme mix (catalog # M0554S), which claims to require only a 5 min incubation at room temperature.
    • Optionally, heat inactivate the reaction by incubating at 65 °C for 20 minutes; not necessary for immediate transformation.
Component Volume (μL) Note
PCR product 2 No cleanup necessary
DpnI (20 U/μL) 2 NEB catalog # R0176S
T4 PNK (10 U/μL) 1 NEB catalog # M0201L (als see this page)
T4 DNA ligase (400 U/μL) 1 NEB catalog # M0202L
10x T4 DNA ligase buffer 1 contains 10 mM ATP; NEB catalog # B0202S
ddH2O 3
  1. Transformation.
    • Because the PCR amplification is geometric, a successful PCR should results in relatively high transformation efficiency.
    • Following this protocol will still give you some colonies with the WT plasmid. As such I usually pick 4 colonies for sequencing.

Gibson assembly mix prep

  • Protocol for making homemade gibson assembly mix (AKA isothermal assebbly reaction mix). Ming previously prepared it, and for many years in the Kortemme Lab everyone was using it.
  • Paper: Gibson, D. G., Young, L., Chuang, R. Y., Venter, J. C., Hutchison, C. A., & Smith, H. O. (2009). Enzymatic assembly of DNA molecules up to several hundred kilobases. Nature methods, 6(5), 343-345. https://www.nature.com/articles/nmeth.1318

5x isothermal assembly reaction buffer (assemble on ice):

  • Reagents:

    From the paper Actually used
    3 ml 1 M Tris-HCI pH 7.5 3 ml 1 M Tris-HCI pH 7.5
    150 ul 2M MgCl2 300 ul 1 M MgCl2
    60 ul 100 mM dGTP 600 ul 10 mM each dNTP
    60 ul 100 mM dCTP
    60 ul 100 mM dTTP
    60 ul 100 mM dATP
    300 ul 1M DTT 300 ul 1M DTT
    1.5 g PEG-8000 1.5 g PEG-8000
    300 ul 100 mM NAD 20 mg NAD 25
    ddH2O to 6 ml ddH2O to 6 ml

  • Prepare 320 ul aliquots (18), label these "5X isotherm buffer", and freeze (leave one to make assembly mix)

isothermal assembly mix

  • To one 320 ul aliquot of buffer, add:

    Volume Reagent
    1.2 ul T5 Exonuclease
    20 ul Phusion polymerase (NOT HOTSTART)
    160 ul Taq ligase
    700 ul ddH2O

  • Prepare 15 ul aliquots (-80) on ice in PCR tubes and store at -20C.

  • These should be good for up to a ~1 year.

RbCl Competent cell prep

Tina shared this protocol for making competent Top10 E. coli cells for plasmid transformation (for cloning). Chris used it in May 2021 and got good transformation efficiency. It is very similar to the McManus Lab Protocol.

Materials

  • TFB1:
    • 100 mM RbCl (MW = 120.92)
    • 50 mM MnCl2, 4xH2O (MW = 197.91)
    • 30 mM potassium acetate (KCH3CO2, MW = 98.14)
    • 10 mM CaCl2, 2xH2O (MW = 147.02)
    • 15% Glycerol
    • adjust to pH 5.8 with 1M acetic acid (0.2%, 1M CH3COOH), do NOT overshoot, filter sterilze, and store at RT but chill to 4C before use
    • NOTE: To avoid precipitation of MnO2 (brown colored solution or precipitate), dissolve everything except MnCl2 in 3/4 of the volume, adjust pH to 6.0 with acetic acid, add MnCl2 (dissolved in water), fill up with water nearly to the required total volume and then further adjust the pH to 5.8 if it is not already there. Avoid adding KOH (or any base) to a solution containing a Mn2+ salt!!
  • TFB2:
    • 10 mM MOPS (MW = 209.3) or PIPES
    • 10 mM RbCl (MW = 120.92)
    • 75 mM CaCl2, 2xH2O (MW = 147.02)
    • 15% Glycerol
    • adjust to pH 6.5 with KOH, do NOT overshoot, filter sterilze, and store at RT but chill to 4C before use
  • LB media and plates

Steps

  1. Culture bacteria
  • Streak/plate bacteria of choice on LB agar plate.
  • Inoculate single colony into 2.5 ml of LB + antibiotics (if used)
  • Incubate overnight in 37C shaker.
  • Subculture the overnight culture 1:100 by inoculating 1ml into 100ml of pre-warmed LB + antibiotics (if used) in a 250 mL erlenmeyer flask.
  • Grow at 37C until OD600 reaches 0.4 to 0.6 (~5h)
  1. Collect and treat bacteria.
  • Cool the culture on ice for 5 min and transfer to a sterile, round-bottomed centrifuge tube
  • After this point, keep everything cold. Work in cold room and pre‐chill all supplies.
  • Pellet the cells by centrifugation 4000 x g, 4C, 5 mins.
  • Discard the supernatant, keep the tube with the pelleted cells on ice.
  • Gently resuspend the cell pellet in ice-cold TFB1 (30 mL for a 100 ml culture). Combine the resuspended cells in one bottle.
  • Incubate on ice 90 minutes at 4C.
  • Pellet the cells by centrifugation 4000 x g, 4C, 5 mins.
  • Discard the supernatant carefully, keep the tube with the pelleted cells on ice.
  • Gently resuspend the cell pellet in 4ml ice-cold TFB2.
  • Incubate on ice 15‐60 mins.
  1. Storage
  • Aliquot 100 ul/tube for storage at -80C. Quick-freeze the tubes in LN2 or a dry ice/isopropanol bath
  • Store at ‐80C in freezer boxes without dividers.
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