Results tagged “protocol” from The SIBYLS Beamline
|Detergent||Category||Concentrations Shot||MW||CMC (mM)||CMC (%)||Type||Aggreg. # *||MW x N (kD)|
|DDMAB||1||0.5x, 1x, 2x||299.5||4.3||0.129||Z|
|n-Hexadecyl-ß-D-maltoside||1||0.5x, 1x, 2x||566.6||0.0006||0.000||N|
|C-HEGA-8||2||0.5x, 1x, 2x||293.3||180.0||5.279||N|
|n-Octyl-ß-D-glucoside||2||0.5x, 1x, 2x||292.4||24.5||0.716||N|
|Nonyl-ß-D-glucoside||2||0.5x, 1x, 2x||306.4||6.5||0.199||N|
|TRITON® X-100||2||0.5x, 1x, 2x||631.0||0.9||0.057||N|
|CHAPS||3||0.5x, 1x, 2x||614.9||8.0||0.492||Z|
|Cymal®-3||3||0.1x, 0.2x, 0.5x, 1x, 3x||466.5||34.5||1.609||N|
|MEGA-8||3||0.1x, 1x, 3x||321.4||79.0||2.539||N|
"category" - group designation based on 1xCMC scattering curve
"N" - non-ionic detergent
"I" - ionic detergent
"Z" - zwitterionic detergent
"Aggreg. #" - monomers per micelle (from CalBiochem)
"MW x N" - estimate of micelle molecular weight
|Below CMC||Above CMC|
|Good for SAXS||Possible with strong protein signal|
|Good for SAXS||Bad for SAXS|
|Possible with strong protein signal||Bad for SAXS|
Here are the recommended concentrations of detergents that may contribute the least amount of background to SAXS studies. Omitted from this list are any detergents that would need to be used at concentrations < 0.1%, based on the above table and the detergent CMCs.
Group I less than 2xCMC
Group II less than 1xCMC (rounded down)
0.2% n-Nonyl-β-D-maltoside (2)
1.2% Zwittergent® 3-10
9.2% Zwittergent® 3-08
Group III less than 0.2xCMC
Detergents can be extremely useful in preparing monodisperse samples for SAXS. However, they can also contribute enormous background scattering, potentially masking the desired protein signal entirely.
We recommend to not using detergents in entire protein preparation. We observed only a few cases (~0.1%) where the detergent ( bellow CMC concentration ) did not affect the protein signal. Basically, there is a high chance that detergent will make your SAXS data useless!!!!
It is crucial, therefore, to choose a detergent that will contribute the least amount of background scattering to your sample while improving sample monodispersity. The data presented here (see links at bottom of page) should be a useful starting point in choosing the right detergent for your sample. Use a detergent only in the case when the protein preparation without detergent was not successful
How we looked for good detergents
We searched for SAXS-compatible detergents by collecting data on 73 detergents in PBS. By subtracting the PBS signal from the detergent signal, we can observe the scattering contribution of detergent alone. We first collected data at 1xCMC (critical micelle concentration) to compare the scattering effects of different detergent types. Next, we investigated the effects of concentration relative to the CMC, collecting data ranging from 0.1xCMC to 3xCMC on a subset of the 73 detergents.
Scattering signal is highly concentration-dependent (below CMC is much better than above CMC), but not all detergents are created equal. Based on the 1xCMC scattering curves, we derived three groups of detergents with scattering properties ranging from essentially no signal (group 1) to large protein-like signal (group 3). The following table provides a rough guide of detergent SAXS-compatibility:
|Below CMC||Above CMC|
|Good for SAXS||Possible with strong protein signal|
|Good for SAXS||Bad for SAXS|
|Possible with strong protein signal||Bad for SAXS|
- 1xCMC may not be a very reliable data point
- Sensitive to pipetting error
- CMC fluctuates with temperature, pH, ionic strength
- Majority of data from single experiment
- Unclear why detergents scatter differently; no obvious trends in MW, micelle size, % concentration, etc.
- Detergents alone may behave differently than detergents in complex with protein
- Some level of subjectivity in "group" designations
- Doesn't address membrane protein applications
What the data provides, however, is a starting point for finding a detergent that improves the monodispersity of your sample without dominating the SAXS signal.
The following table include links to scattering curves of each concentration of detergent we analyzed, with Glucose Isomerase plotted as a reference protein curve.
A sortable Table of detergents.
List of best-bet detergents
The detector can be brought up to essentially room temperature (in order to gracefully shut it down or do other work on it) or brought down to -40°C (to minimize detector noise for data collection) using the “Quantum Console” program on the detector computers.
This figure shows a fairly typical screen image on the detector computers.
In the image shown, the “REMOTE Detector OP” program is already running (as a DOS window). If this program is not running, it needs to be run before the Quantum Console can control the detector modules. The shortcut to the program is highlighted by one of the white circles.
Once the “REMOTE Detector OP” program, hereafter the detector program, is running on all nine modules, you can control the detector temperature by running the Quantum Console program on any one of the detector computers. The shortcut for this program is highlighted by the other white circle.
When the Quantum Console starts, it will open a window that looks something like this:
Before you can do anything else, you need to click on “Connect to Detector Processes.” This will create a row of status reports and allow you to click on “ENABLE Temperature Control”, resulting in a window something like this one:
This window actually shows a temperature-change operation that is already under way, but all the essential parts are visible.
If all is as it should be, setting the temperature is fairly simple. To cool the detector down for operation, click “Ramp to Cold Operating Temp”. To warm it up, click on “Warm up Detector to +10°C”. Either one will gradually bring the detector to the desired temperature. Shifting by the whole range from +10°C to -45°C, in either direction, generally takes about an hour.
Unfortunately, it is occasionally necessary to restart the ADSC detector along with the associated software. This will summarize how that is done. We will assume that everything is up and running at the beginning of the process; starting when some component or other is down is essentially the same.
We’ll assume that this is only a short interruption and that we don’t have to worry about the detector vacuum.
Begin by bringing the detector to room temperature (or +10C) as described in “Detector Temperature Settings.”
Once the detector is warmed up, it can be shut down at the power switches —- it is not necessary to shut down the detector processes or other software first.
Turn the detector power switches (the green switches highlighted by the arrow) off from top to bottom. The main power switches should be turned on starting with the bottom power module, then the middle module, then the top module. The order does matter.
If you don’t have the “REMOTE Detector OP” processes running yet, this is the time to start them. Double click on the “REMOTE Detector OP” shortcut, and you should see a DOS window appear:
Even if you already had these processes running when you restarted the detector, they still must be sent a reset signal. We generally do this from a script. The script makes up part of /programs/beamline/nuke, so it can be found there, but for completeness, here is the one we’ve been using:
foreach module ( detector0 detector1 detector2 detector3 detector4 detector5 detector6 detector7 detector8 ) # send the reset signal echo -n "$module restart " echo "restart" | sock_exchange.tcl $module 8038 1 echo "" end
Entering that script at a shell prompt at any of the BL12.3.1 Linux consoles should reset the whole group of detectors. (Since the BLU-ICE/DCSS control programs run on dataserver, that is a good place to run the script.) This reset signal will cause lots of activity on the “REMOTE Detector OP” windows:
Focus Beam at Beamstop with Experimental Table Between PX and SAXS
Check Helium tank outside of hutch to make sure it is full and that it is flowing into the shutter box. It is important to purge all air from the shutterbox as the ion-guage needs to report a consistent value.
Turn Helium on as early as possible to assure shutterbox is purged
In a terminal window turn off the feedback with the following command:
Switch KVM to the Beamline computer, and launch "Shortcut to BL Control Main".
Click the white arrow in upper left of main window. This will open the "Beamline 12.3.1 Beamline Control System" window.
Open bothe the Motor Debugger and Motor Monitor modules from the "Motors" pulldown menu.
Turn on autoscaling under the "Amplifiers" pulldown main menu. (why?)
M2 Bend Up to 244000, and click move button.
M2 Bend Down to 267000, and click move button.
Mono eV" to 10000.
Remove the safety pin and use the hand crank to move the table to the mid point. There is a piece of red tape marked in pen "YAG" indicating the proper midpoint position of the table.
Attach the BNC cable to the camera that monitors the YAG prism, and focus the camera (this step will hopefully be unnecessary once a more permanent camera position is established.
Close the hutch.
Open the main shutter. You should see the direct beam hitting the YAG prism at this point.
tune rocking curve" and click either the step ⬆ or the step ⬇ button once to initialize the optimization procedure. (this automagically adjusts Theta2)
M2 Tilt" and click the home button to reset all values to 0.000
Move M2 Tilt to 12000.
Some PX users will have adjusted the Slits1 to make a very tight beam so they should checked and backed off if necessary. Aperture Line 1 and Aperture Line 11 values should be decreased several unit values.
If you lose the beam jog
M2 Tilt in 100 unit increments (this should move the beam up and down on the video monitor).
Chi2" (value should be ~0.492) jog Chi2 in 0.01 increments. (this should move the beam left and right on the video monitor). Because Chi2 is adjusting the focus of the beam changes to this value will drastically alter the shape of the beam.
Sometimes the table will need to be moved slightly in order to position the beam in the middle of the YAG. Additionally the shutter control cable may sometimes get in the way so it must be unplugged from the shutterbox. Turn off power on shutter control box and unplug cable.
Close hutch and turn off hutch light.
Open main shutter and make sure video monitor is turned on.
Optimize the shape and size of the beam. You want it as round and small as possible. Make small adjustments to both Chi2 and M2 Tilt.
Move Experimental Table to SAXS Position
Move the table all the way into SAXS mode using the hand crank. There is a digital dial attached to the air table's sub-frame. Crank the table until this value reads zero.
Re-attach the shutter control cable to the shutter box and turn on the power to the shutter control box. Press the small red button on the shutter control box to really turn it back on.
Close hutch and open main shutter.
Optimize Slits 1, Slits 2, and Slits3 - itertatively
Make sure that the video signal going to the beamline computer is displaying the Beam Position Monitor (BPM) video-feed from the camera that points into the shutter box.
Turn on feedback with the following command:
The point here is to move slits1 (Aperture guys) and slits2 (SAXS aperture guys) and slits3 (Guard Slits guys) back in as far as possible without actually clipping the beam.
The beam on the back of the shutter may be a bit rough on the bottom edge as a result of nicking slits2 and may look like this:
Jog M2 Tilt to move the whole beam slightly up.
Make sure the High Voltage control unit that feeds the ion guage in the shutterbox has not been tripped. If it has been tripped then reset and increase voltage to as close to 300V as possible.
Close all four slits1 blades as far as possible without clipping beam.
Aperture Line 1moves in from the right
Aperture Line 11moves in from the left
Aperture Uppermoves in from the top... duh
Aperture Lowermoves in from the bottom... duh
Close all four slits2 blades as far as possible without clipping beam.
SAXS Aperture Line 1moves in from the right
SAXS Aperture Line 11moves in from the left
SAXS Aperture Uppermoves in from the bottom... huh?
SAXS Aperture Lowermoves in from the top... huh?
Insert all filters into direct beam and take a 1sec shot on the MAR CCD.
Close all four slits3 blades as far as possible without clipping beam.
Guard Slits 1more + values to close blade down. (Left side)
Guard Slits 11more - values to close blade down. (Right side)
Guard Slits Uppermore + values to close blade down.
Guard Slits Lowermore - values to close blade down.
Reduce Reflections from Slits3 and adjust beamstop
Take a 1 sec shot on MAR CCD. It will look something like this:
Back off the Guard Slits Upper and Guard Slits Lower until the vertical streaking is eliminated. (streaking toward the top of the screen is caused by Guard Slits Lower being too far in, and vice-a versa for streaking downwards)
Back off the Guard Slits 1 and Guard Slits 11 until the horizontal streaking is eliminated. (this is usually less of a problem, but again streaking to the left is caused by x-rays reflecting off of Guard Slits 11, and vice-a-versa for streaking to the right)
Adjust Endstop y to move the beamstop up and down.
Adjust Endstop x to move the beamstop left and right.
Center the beamstop by observing a line integration tool drawn though the beamstop. The idea is to get a symmetrical background scatter aboce and below the beamstop shadow.
Define Beam for Users
Enter hutch. Manually open the experimental shutter. Insert CCD shield. Insert sample cell with fluorescent paper.
Close hutch. Open main shutter and define the bounding box for the beam on the video monitor.
It may be necessary to move the sample up or down (Sample x and Sample y) so that the beam enters the sample cell in the center.
Enter hutch. Close experimental shutter. Remove CCD shield.
You are now ready to CRUSH!!!!!
- Macromolecule sample and exact buffer
- The sample cell takes 15 ul. 20 ul is safer when using the robot.
- Minimum concentration is 1 mg/ml
- Maximal concentration is 10 mg/ml
- Identical buffer required (>5ml recommended)
Please download and fill out this mandatory shipping form before sending your samples. It should be included with all samples sent to the SIBYLS beamline for data collection.
Send samples to:
Lawrence Berkeley Lab 1 Cyclotron Road MS 6R2100 Berkeley, CA 94720 ATTN: Kathryn Burnett / 12.3.1 510-685-5755
Send an email to Kathryn Burnett with federal Express shipping information so she knows the sample is coming and so she can track the sample online.
RECOMMENDATIONS for Sample preparation ( for routine analysis and first-time users )
We recommend not using detergents in entire protein preparation. We observed only a few cases (~1%) where the detergent ( below CMC concentration ) did not affect the protein signal.
The most common problem at the beamline is aggregation in the sample. Since larger particles scatter X-rays more strongly than small particles (albeit less than visible light scattering), aggregation will bias the results. We strongly recommend doing either DLS, native gel, or gel filtration (best). If your sample has a tendency to aggregate over time, it is possible to prepare the sample at dilute concentrate and then concentrate just prior to data collection.
The higher the concentration, the better the signal. However, there's a balance between problems with aggregation/oligomerization at the higher concentrations, unless the macromolecule is well-behaved. We generally recommend 1-5 mg/ml and doing a concentration series. If we are given 50 ul, we can do serial dilutions. Homo-oligomers can give okay signal down to 1 mg/ml. Ideally data should be collected on at least three different concentration of the macomolecule in the range 1-10 mg/ml to identify any concentration dependent behavior. Aggregation precludes data analyses. Concentration determination: OD280 is considered the most accurate although buffer subtraction is very important (oxidized DTT has an absorbance at 280). We have a nanodrop at the beamline (2 ul sample vol), and we recommend taking conc. right after removal from dialysis. Please verify that your conc. is correct--overestimation is a common problem.
96 well plate sample format. Our beamline operates with a pipetting robot, which works with 96 well plates. The samples can be shipped in fast frozen state in the single concentration with minimal volume 24uL .
We strongly recommend dialyzing sample. The difference between the scatter of the macromolecule and buffer is so low, that simply making up the "equivalent" buffer is not sufficient to get accurate subtraction. To retain sample volume, we have found that the Hampton dialysis buttons (eg. 30-50 ul size) are ideal at keeping the volume constant. If you haven't used the buttons before, practice with saran wrap. Hampton has a nice set of instructions--many beginners fine the golf tee to make the difference. To remove the sample, we use a Hamilton syringe with a blunt needle, pierce the dialysis membrane, and remove the sample. We usually lose only a couple ul. The dialysis buttons fit well into 50 ml tubes--I have crammed in 7-8 in the same tube. The 50 ul volume allows us to do serial 1:1 dilutions. The buffer in the concentrator flow through is also good for subtraction (depending on your buffer components).
Salt increases the background, but we've gotten good signal with up to 1 M salt. Concentration of the macromolecule has more of an impact on signal than the buffer, so if the sample is monodisperse in high salt, put it in high salt. 1-5% glycerol cuts down quite a bit on radiation damage, although we've also seen oligomerization induced by addition of glycerol. If you have the quantity available, trying several different buffers is recommended. For a start, ideal buffer might contain 100-200 mM salt with 5% glycerol. Please send at least 15 mL of buffer with your sample.
This happens, maybe 1 in 10 samples we observe changes in the SAXS scattering curve due to radiation damage. We've found that complexes do seem more stable, so if you have a complex, then throw that in as well. Glycerol is a pretty good radical scavenger, so 5% glycerol is good.
The beamline is equipped with Peltier that can vary the temperature from 0 deg to 50 deg.
IMPORTANT INFO TO INCLUDE FOR COLLABORATIONS:
If you are sending in your sample for data collection as a collaborative effort with a beamline scientist, please
- 1. Label tubes uniquely so that when they are stuck in the frig with many other tubes, they don't get lost. Recommend your initials with a number (eg st1) and date of sample.
- a. Sample
- b. Corresponding buffer
- 2. Send information on the sample that will help during data collection:
- a. MW and oligomerization state, if known
- b. Number of residues
- c. Extinction coefficient 280
- d. Buffer components
- e. Sample concentration done on concentrated sample and what method used.
DATA COLLECTION BASICS:
Sample cell has flat Mica windows. We generally keep our Pilatus 2M detector at 1.5 m distance.
We have developed a time slicing mode for collection. Each sample is collected multiple times with the same exposure length. We generally collect every .3 seconds for a total of 10 seconds resulting in 30 frames per sample (though there is some variation depending on beam optimization).
The time of the short exposure is selected to minimize detector overloads near beam stop but to provide accurate measurement close to the beam stop. The long exposure is generally ten-fold longer to accurately measure high angle data.
We can change the X-ray wavelength to optimize for sample size, although 11 to 13 KeV is generally fine for most samples under 40-150 kD MW. Qmin should be collected at an angle < pi/dmax. Detector parameters however must be calculated for each wavelength (we use silver behamate), so if you want to change the wavelength, please advise the beamline scientist.
1. Circular integration, normalization, and subtraction of sample and buffer image files. Usage --> ogreNew sampleimg bufferimg
ogreNew *005 *004
[subtracts image4 (buffer) from image5 (sample) and generates I vs S scattering curve]
2. We use xmgrace to visualize the scattering curves.
3. Preliminary analysis using PRIMUS and Gnom during data collection (from Svergun's group) is extremely helpful in determining the quality of your data. If your data is not good, subsequent analysis will be inaccurate.
PRIMUS-guinier analysis indicates aggregated sample (increasing and non-linear slope towards I0). I0 estimate also good for estimating molecular weight using known standards (MW is linear with I0/conc).
PRIMUS-porod analysis gives rough estimate of molecular weight (porod volume*1.2/2 ~ molecular weight).
PRIMUS--SASPLOT (I*S^2 vs S) gives indication of folded nature of molecule. For example, well folded proteins show parabolic curve that returns to baseline. Unfolded or random coil molecules will increase with angle. Please note that too dilute sample and poor signal at high S or inaccurate background subtraction will also give this result. GNOM--p(r) analysis. Aggregated sample will have unstable Rg and p(r) curve will have small bump close to Dmax.
A new review on macromolecular SAXS has been published in the Quarterly Reviews in Biophysics by Putnam, C.D., Hammel, M., Hura, G.L., and Tainer, J.A.
“This six part review addresses both theoretical and practical concepts, concerns and considerations for using these techniques in conjunction with computational methods to productively combine solution scattering data with high-resolution structures.”
The review provides an extensive and up-to-date review on the application of small angle X-ray scattering is available for download.