Prepare for your visit

Batch SAXS: users must provide a matched buffer solution for background subtraction.
This should be the exact same buffer used in the original sample preparation if possible. Change buffer (by centrifugal concentrator, dialysis, or SEC) if you don't know the original composition exactly. Prepare plenty of extra buffer for sample dilutions and for rinsing sample cells (bring at least 10 ml if you can).

Protein solutions should be monodisperse and concentrated to at least 5-10 mg/ml if possible without aggregation. It is a good idea to bring at least some unconcentrated sample to test for concentration-induced aggregation. You should check monodipsersity using dynamic light scattering (DLS), if available. Purifying your sample on a size-exclusion column is highly desirable. Even with pure proteins it may be necessary to experiment to find the best conditions under which the protein is monodisperse and well folded. Sometimes proteins will aggregate rapidly after purification and must be purified on site. Protein samples have been frozen successfully for shipment, but users should not assume their proteins will necessarily freeze well.

A small amount of glycerol (1-5 %) is advisable for protection from radiation damage, but more than 15% will degrade the strength of your scattering signal and should be avoided. In general, high salt (> 1M) should be avoided for the same reason. Preservatives like DTT and TCEP are known to reduce radiation damage in small amounts and are therefore recommended.

We strongly suggest that you measure your concentration as accurately as possible! Molecular weight estimates based on known concentration are valuable indicators of sample monodipsersity, oligomeric state, and can help diagnose problems with mixtures. A280 UV sample measurements are possible at the beamline, if needed.

Individual exposures can be done with sample volumes as small as 20 microliters, if loaded by hand. But for easy and reliable sample loading, we recommend using the sample loading robot with at least 30 microliters if possible. Larger sample volumes ensure easy and rapid positioning of sample in the beam and a generous oscillation size which will eliminate potential radiation damage. Since BioSAXS requires multiple dilutions (at least on some representative samples), it is advisable to bring more than the minimum volume. A sample size of 50 microliters, for example, would allow just enough to prepare a minimum series of 3 dilutions of 30 microliters each: full strength (30 ul sample), 66% (20 ul sample + 10 ul buffer), and 33% (10 ul sample + 20 ul buffer). More dilutions are desirable, if possible. Many users prefer to prepare dilutions by halving: 1.0, 0.5, 0.25 0.125 etc.

With a flow cell, the more protein you bring, the better the signal will be. For lysozyme-sized proteins (14 kDa), don't expect to get much usable signal below 1 mg/ml. For larger proteins, the low concentration limit will improve. Glucose isomerase, for example, (MWt = 173 kDa) gives good data at 0.3 mg/ml and is essentially at the infinite dilution limit.

Solutions that are too concentrated exhibit concentration-related distortions of the small-angle part of the scattering curve. You can see this in lysozyme stronger than 10 mg/ml and Glucose Isomerase stronger than 0.5 mg/ml.

It is advisable to collect data at several (at least 3) different concentrations and extrapolate to infinite dilution if necessary. Alternatively, you can combine a dilute curve (small-angle part) with a concentrated curve (wide-angle part). Concentration does not effect the wide angle part of the scattering curve.

The concentration of protein necessary to get a good signal in Batch BioSAXS depends inversely on molecular weight. For small proteins like lysozyme collected on S7A station using typical exposure times, 4 mg/ml will usually produce a profile that is dilute enough to avoid interparticle interactions, but strong enough to give a good low-noise Guinier plot with accurate radius of gyration. Larger proteins like glucose isomerase (173 kDa) need only reach 0.3 mg/ml to give the same strength of signal in the low-angle region. A good rule for "dilute" sample concentration is 60/(molecular weight in kDa) = concentration (mg/ml). Best to bring more concentrated sample (x5) so you can create a concentration series. Generally, SEC-SAXS requires 100 microliters of  sample at 5x60/(molecular weight) since chromatography tends to dilute samples.

Short answer:  for Batch SAXS, expect to measure approximately 80 samples in the first 24 hours. Experienced users may collect far more. As of Sping 2023, we are in process of upgrading our sample-handling robot. For inline chromatography (SEC-SAXS) using the short 5/150 column (good for cleanup of samples, but not for high-resolution separations), expect measure at least 25 samples in 24 hours, if you have no buffer changes. For every buffer change, subtract 2 samples. High-resolution SEC-SAXS separations using the 10/300 column take roughly twice as long. Actual performance will vary widely. With experience, and with your group working as a team, these numbers could be significantly higher.

Actual time for data collection is hard to estimate because there are so many variables. With our new capillary-flow cell, we can now take much longer exposures than before without radiation damage. This can dramatically improve data quality, but it also will reduce the number of samples that can be examined in the time available. Currently we are recommending at least ten 1 second exposures per sample, so the total exposure would be 10 seconds. Each protein you examine will include a buffer exposure of equal length and at least 3 concentrations. You may use the same buffer profile for multiple proteins, if you like. But, it is *very* wise to re-take the buffer periodically. You may wish to expose dilute solutions for longer using larger sample volumes. You should also factor in sample cell rinse and dry time between different proteins, which will add ~2.5 min. Spinning samples prior to data collection requires 5-10 min at 14,000 rpm, but sample spinning and dilution can be done simultaneously with data collection.

• aggregation (most common)
• molecule too large (beamline can't reach low enough q)
• sample is a mixture (common with complexes)¹
• sample too dilute
• radiation damage
• denaturation (rare)
• buffer mismatch
• contrast problems (weak signal due to buffer composition)
• heterogeneous sample (protein-DNA-lipid complexes)²

¹Generally, solutions must be pure and monodisperse for standard SAXS analysis to work. If you are hoping to see a conformational change when you add a component like a small ligand or other binding partner to the protein sample, be careful to maintain exactly matching buffer. This may mean changing buffer again using a centrifugal concentrator, dialysis, or a SEC run. If conversion of your protein to the new state is incomplete, you may need to re-purify. Mixtures can be treated with BioSAXS, but this is an advanced topic and additional information and multiple experiments may be required to understand the data.

²Mixed complexes of protein, DNA, and/or lipids can cause problems with certain SAXS calculations.

MacCHESS provides several protein standards for use in your SAXS experiments. It is important to run at least one standard so that you have a way to estimate molecular mass, but also a way of being confident that the beamline is running properly. Standards also serve as a reminder of what good monodisperse sample *should* look like.

Lysozyme Buffer:
40 mM NaOAc pH 4.0
150 mM NaCl
1% glycerol v/v

Glucose Isomerase Buffer:
10 mM HEPES pH 7.0
1 mM MgCl2

Protein concentrations vary from run to run, but will be approximately 4 mg/ml for lysozyme and 0.4 mg/ml for glucose isomerase. A more complete list of standards can be found in:

Mylonas, E., and D. I. Svergun. 2007. Accuracy of molecular mass determination of proteins in solution by small-angle X-ray scattering. J. Appl. Cryst. 40:S245-S249.

Kozak, M. 2005. Glucose isomerase from Streptomyces rubiginosus - potential molecular weight standard for small-angle X-ray scattering. J. Appl. Cryst. 38:555-558.

MacCHESS staff can teach you how to create scattering profiles, evaluate data quality, and compute R_g on site. But, just like protein crystallography, there is a lot to learn … much more than you can absorb in one sitting. We strongly recommend that new users take a training course, if possible. Such 1-2 day courses are offered by a number of synchrotron sources. MacCHESS periodically offers an introductory workshop entitled BioSAXS Essentials. To receive notification of the next scheduled course offering, please visit this link.

Yes, Batch BioSAXS can be performed at any temperature from 4ºC to 80ºC. SEC-SAXS only supports 4ºC operation since the chromatograph is situated in a refrigerator. HP-SAXS can be performed from 4ºC to 80ºC and HP-SEC-SAXS can be performed from 4ºC to 50ºC.

We have calibrated the actual sample temperature using a micro thermocouple and provide a correction table for setting the chiller for normal BioSAXS operation. Notes for the provided spreadsheet program (courtesy of Matt McLeod): 

Air is just the air temperature 
Liquid is just liquid in the capillary - not oscillating and not incubated in the block
Oscillation is when the plug is oscillating - this takes less than a minute to get to temperature.  

The sample as it moves into the capillary heats significantly and just needs a little more time to get fully equilibrated. I then did a 5 and 10 minute incubation in the block, followed by moving into capillary for temperature measurement.  If not oscillating the temperature steadily goes down to get "liquid" temperature.

CHESS_7A1_temperature.xls

Anoxic SAXS (anSAXS) allows samples to be prepared within an anoxic chamber (<30 ppm O₂), which is directly connected to the X-ray sample cell so that samples and buffers are kept fully anoxic throughout an experiment. Users can perform both batch and SEC-coupled SAXS, with the ability to spin down samples in a refrigerated centrifuge and collect UV-vis spectra within the anoxic environment. SEC-anSAXS has in-line full UV-vis range spectroscopy capability, so time series on absorbance from 200-800 nm can be measured throughout an experiment. A full spectrum can also be collected at any point during the sample elution.

AnSAXS samples can be shipped to CHESS or brought to the beamline frozen, being allowed to thaw only once they are in the anoxic chamber. Sample and buffer considerations are the same as with conventional batch SAXS and SEC-SAXS. If planning to prepare buffer on site, house nitrogen is available for buffer sparging to reduce dissolved oxygen.

Sample concentrations for anSAXS experiments are like that of conventional batch SAXS and SEC-SAXS, however components that will be used during the experiment such as centrifugal concentrators, buffer exchange columns, or other forms of non-standard tubing must be arranged in advance to allow time for gas equilibration within the chamber.

Temperature control is different for anSAXS than it is for conventional SEC-SAXS due to the chromatography column being in the anoxic chamber which is currently not temperature controlled. Heat accumulation from mechanical components within the chamber has been minimized to a great extent, but the temperature within the chamber is often greater than the surrounding room. For batch anSAXS, the sample can remain on a cold block and at 4°C within the refrigerated centrifuge until loading, when it is transferred to the X-ray sample cell which can be temperature controlled.

When you use the spec command "rgseries <number of snapshots> <exposure time per snapshot>", the X-ray shutter will open and the detector will take "number of snapshots" images with no delay between images.  Each image will accumulate photons for "exposure time per snapshot" seconds. When finished, the shutter will close. For batch SAXS experiments, the sample will have been exposed to X-rays for a total of (number of snapshots)*(time per snapshot) seconds. Longer exposure time or more snapshots will increase the total dose. This can improve the signal strength, but also risk more X-ray damage. Finding the right total dose is a matter of experimentation, but your beamline scientist will start you with recommended settings based on common proteins.

For SEC-SAXS, use rgseries, but simply give a large number of images. For example: rgseries 1500 2. Changing the exposure time, 2 seconds in this case, does not change the dose because the shutter is open all the time and the sample is flowing past the beam. Using 1 second exposures will give you twice the number of points on your chromatogram, but the individual exposures will have more noise. The only way to change the actual dose is to change the attenuation of the X-ray beam or to change the chromatographic flow rate. The beamline scientist will show you how to do this, if needed.

When RAW reads a detector image, it applies a normalization factor based on the integrated transmitted intensity. As a consequence, scattering profiles taken at different exposure times and different attenuations will actually overlay (reasonably well). Normalization is applied because synchrotron beams can fluctuate in intensity over time (during topoffs, for example). So normalization allows you to compensate for slight differences in sample dose. It is not intended to compensate for large changes that might result from intensional attenuation. So, it is always recommended to subtract like exposures with like exposures, when possible.

Finally, RAW usually applies a scale factor to place the scattering profile on an absolute scale. As a consequence of how normalization is performed, scattering profiles will always be on an absolute scale regardless of exposure time or incident beam intensity.

It is important at the start of any visit to verify all the above settings in the RAW software and save the appropriate .cfg file containing those settings.