Prepare for your visit
How to prepare samples
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.
What sample concentrations do I need?
The concentration of protein necessary to get a good signal in BioSAXS depends inversely on molecular weight. For small proteins like lysozyme collected on F2 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.
How many samples should I bring?
Short answer: expect to measure approximately 80 samples the first 24 h. Actual performance will vary.
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 2 second exposures per sample, so the total exposure would be 20 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 wise to re-take the buffer periodically after each re-filling of the synchrotron storage ring. 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 min. Spinning samples prior to data collection requires 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.
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.
How long does it take to learn to process SAXS data?
MacCHESS staff can teach you how to create scattering profiles, evaluate data quality, and compute Rg 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.