On Fri, 16 Feb 2001 12:38:21 -0800, david.mccready@pnl.gov (Dave McCready) wrote:
------------------ Poor Man's "Anaerobic" Specimen Mounts for XRPD --------------------------
Below are two inexpensive means of handling air-sensitive specimens for XRPD analysis. Both methods assume a standard powder diffractometer (no special sample stage) and are based on the use of a front-loading cavity-type holder that is air-tight except for the normally open front surface.
First, an important disclaimer. As far as I am concerned neither method outlined below is subject to debate here. If you have something to add, great. But just like your mother told you, if you have nothing nice to say then don't.
----------- KAPTON Tape -----------
Prepare and mount specimen per usual (e.g., in Ar-filled glove box or glove bag). Cover mounted specimen with KAPTON tape, making sure the tape is securely fixed around edges of cavity. Mounted specimen covered with KAPTON tape can now be safely exposed to air for several days without fear of oxidation, etc.
Of course, one "sees" the KAPTON tape in the diffractogram (it produces a high amorphous background), but with modern methods of data collection and processing this is not a significant problem. (One exception would be cases where determination of the amorphous content of the sample is a goal of analysis.)
A commercial US source for KAPTON tape is CS Hyde Company (800.461.4161 or http://www.cshyde.com/kapton.htm). Cost is just a few to several dollars per roll, and the company accepts small orders. I use their #18-1S (0.003 inch thick). In that weight, KAPTON is fairly transparent to the beam.
-------- GLYCEROL --------
Most powders are miscible in glycerol, but will not react with it. Use a thick (i.e., powder-rich) suspension. Flat specimen mounts are a real challenge here, but the sample will again be safe from oxidation for at least a few days. Method works best with T-T machines (where the glycerol will not run out of the holder), but it can also be used on T-2T types up to moderately high angles. Again, this method produces a high background.
----- OTHER -----
Off topic, but another use for KAPTON tape is fixing heat tapes to UHV chambers for bakeout. Facilitates more intimate contact of heat tape to chamber, which in turn promotes more even bakeout. KAPTON itself will readily withstand the high temperature. The adhesive will eventually dry out, but it works better than nothing.
Date: Wed, 28 Feb 2001 18:39:52 -0800
From: "McCready, David E" <david.mccready@pnl.gov>
Subject: RE: Poor Man's "Anaerobic" Specimen Mounts for XRPD Message-id: <29E6E93D92576F4DB85FE89FF2C23F440D3C96@pnlmse03.pnl.gov>
MIME-version: 1.0
If using KAPTON tape, it is important to make sure it is pressed firmly around the edges of the cavity in your sample holder. This will ensure a good seal against the intrusion of air. This was all I meant by "securely" fixing the tape to the sample holder.
Also, I find it convenient to cut several pieces of tape, fold one edge of each piece over to form a non-adhesive tab, and temporarily attach the pieces of tape to a glass slide before putting the whole works inside the glovebox. It is easier and safer to cut the tape outside the box. Folding over one edge makes the pieces of tape easier to handle (especially with big thick gloves on). Sticking them on a clean slide gives you a sort of custom tape "dispenser" you can then use in the glovebox. Having several pieces of pre-cut tape handy allows for quick recovery from mistakes (e.g., tape gets stuck to the glove or something). I also cut the tape in relatively long pieces and make the tab fairly large. Both facilitate handling the tape with gloves.
When I use the KAPTON tape to cover a powder specimen, I hold it by the tab and start the tape onto the holder some distance away from the cavity. Then I sort of press/roll the tape down over the cavity using a small but sturdy glass vial (e.g., a standard 20 ml scintillation vial). At the same time, I maintain some tension on the tape by pulling the tab in a direction upwards and away from the cavity (in the direction I roll the vial). Kind of hard to describe, but if done slowly, this results in a nice, flat KAPTON cover for the specimen.
As for making sure the tape is absolutely flat, I would not worry too much about that. It does not have to be "perfect" to work. You can prove this by expt. Take a slide and stick a piece of KAPTON tape or film to it making sure it is "perfectly" flat, then scan it. Take another slide and stick a piece of KAPTON on it, purposely bunch up or wrinkle the film/tape a little bit, then scan that. The data will be very similar assuming the beamspot covers more or less the same area of KAPTON in both cases (probably best to have the beam entirely on KAPTON in ~all~ cases if possible).
BTW, I like your idea of using KAPTON film and vacuum grease. Thanks for mentioning it in your e-mail. I will have to try it sometime.
Good luck with your work, and please let me know if I can be of further assistance.
Thanks,
Dave McCready
Battelle EMSL
Date: Thu, 22 Mar 2001 18:33:37 -0800
From: "McCready, David E" <david.mccready@pnl.gov>
Subject: RE: Poor Man's "Anaerobic" Specimen Mounts for XRPD
My own sample mounting technique generally results in what I would call a very "clean" specimen. Tools involved include a single-edge razor blade and the type of microscope slide that is frosted on one end. In addition, I strive to have very finely ground samples whenever possible (which is usually, but not always, the case).
The typical result is a flat, dense specimen mount which does not have any excess sample material around the cavity. The KAPTON tape lays down easily with no leaks to air (again, rolling the tape down with something like a 20 ml glass scintillation vial helps).
FWIW, I think preparing "good" XRPD specimen mounts mainly takes practice. At least that's how I learned (I've done in excess of 10,000 XRPD analyses, so I have some practice at the job). Even so, some powders can be very difficult to mount properly.
In any case, I posted an outline of my "standard" XRPD specimen mounting technique sometime ago. For the "anaerobic" approach, one merely adds the KAPTON tape. I also saved a TXT file of that post, and a copy is pasted below my signature. Let me know if you have any questions about it.
Cheers,
Dave McCready
Environmental Molecular Sciences Laboratory
http://www.emsl.pnl.gov
PS: If stuck with the plexi holders, I would recommend using a "Q-TIP" to wipe away excess powder from around the cavity. (a razor blade would very likely damage a plexi holder).
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(Posted to sci.techniques.xtallograpy on 09/09/1999)
As evidenced by recent publications, there appears to be considerable interest in techniques for the preparation of specimens for X-ray powder diffraction analysis. As such, I have decided to throw my 2 cents in here.
First, I will mention my preference in specimen holders. I almost exclusively use the off-axis quartz plates produced by the Gem Dugout in State College, PA (Phone/FAX 814.238.4069). These are high-quality, durable specimen holders, which yield low background. Rather expensive perhaps, but well worth the cost. For most applications, the standard, front-loading shallow cavity versions of these plates are ideal.
Proper X-ray powder diffraction specimens are, of course, flat, densely packed, very fine-grained powders. This is a fairly short order, but I have noticed many people have difficulty accomplishing the task. In particular, achieving high packing density can be a challenge.
The usual approach is to press powder into a cavity using an ordinary microscope slide. However, many materials will agglomerate and pile up upon compression. This tendency interferes with one's ability to also make the specimen mount flat.
My solution has been to use _frosted_ microscope slides.
By frosted, I mean slides that are sandblasted to provide a surface for marking the slide with a pen (not slides that are _painted_ for the same purpose).
I favor slides that are frosted on one end and only on one side. Full-frosted cytology slides might also work, but I have not yet tried them.
Anyway, these partially frosted slides work (for me, anyway) as follows:
------ STEP 1 ------
A small amount of powder approximating (or, ideally, slightly exceeding) the volume of the cavity is placed in the holder.
The _smooth_ portion of the microscope slide is first used to lightly compress the powder into the cavity.
Excess sample material is then scraped away from just the edges of the cavity using a razor blade.
This leaves a slightly elevated specimen surface.
------ STEP 2 ------
Now, the frosted end of the slide is brought into use.
Placing the frosted end of the slide atop the elevated specimen surface, one can drive the remaining excess sample material into the cavity using moderate-to-considerable downward force combined with twisting motion.
Whatever additional excess sample material might then be present appears as a kind of "halo," which will be visible through the slide around the specimen cavity.
If no halo appears, the cavity is not yet "full" (densely packed).
------ STEP 3 ------
A small amount of sample material can be picked up on the corner of a razor blade and deposited on or near the center of the cavity.
Then, the frosted portion of the slide is again used to work this additional sample material into the cavity.
If no halo appears around the cavity, this process is repeated.
Once the halo appears, the cavity is full.
Any remaining excess sample material on the surface of the holder around the cavity can then be (carefully) scraped away with the razor.
Now one has a densely packed, flat specimen mount.
However, its surface is very rough from contact with the frosted microscope slide.
------ STEP 4 ------
As a final step, the surface of the specimen mount can be smoothed using the plain, smooth surface of the microscope slide.
Inspection of the specimen should then reveal a very smooth study surface.
Pits in the specimen surface indicate that the cavity can accept additional sample material. (In that case, return to Step 3.)
-------------- Additional Tip --------------
Certain microscope slides will have frosted surfaces that are a bit too rough for this application. Small particles of glass will be peeled off the slide and mixed into the specimen if very much force is applied in Step 3.
The frosting on such slides can be toned down a bit (smoothed) using a Scotchbrite pad and a little water.
This problem will also occur if the sample material is both coarse (large-grained) and harder than glass. In that case, grind the sample to a finer consistency.
------- Summary -------
Specimens prepared for X-ray powder diffraction as outlined above generally yield very good, reproducible patterns. With practice, such specimens can be prepared in a very short time. Only common and inexpensive tools are required.
The limitations of front-loading specimen mounts are part of the package, but the supposed negative effects of such specimen mounts are, in my opinion, overrated. Specimens prepared by this method will certainly suffice for the vast majority of X-ray powder diffraction analyses.
I hope this information is received in the spirit it is provided, which is that it be of some general interest and utility.
Regards, David E. McCready
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