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If you’ve been following along in the lab, at this point you should have a nice large quantity of functionalized resin sitting in a large-scale peptide synthesizer.  It’s time for the “split” part of our synthesis.

Rinse the beads with dichloromethane, then dry them thoroughly with a stream of air (this takes twenty minutes or so).  Transfer the beads to a pre-weighed five dram vial, a weigh boat, or just our favourite piece of glassware and get their mass.  In an ideal world this will correlate perfectly with the stated loading capacity of the beads and the weight of your peptide chain, but that isn’t critical.  The important thing is to calculate the moles/gram value of your functionalized resin, with a simple equation.

             From there you can calculate the amount of resin required for each of your fragments.  As an alternative path you can also simply divvy the beads into equal fractions, ie. 1/10ths.  This makes weighing all the different coupling agents in the next step much easier, though you’re going to have different a yieldtheory for each compound at the end.

Whichever path you take, the next step is to transfer your beads to some 10mL syringes.  My personal preference is the National Scientific all plastic variety, though it shouldn’t really matter so long as the syringes are chemical resistant.  Cap the syringes with a 22G needle and a small kimwipe plug and you’re good to go, for only a fraction of the price of an automated synthesizer.

Because the beads have been fully dried you’re going to need to resoak them before effective coupling can occur.  So, draw up some DMF through the needle, taking air for the last few millilitres (the bubble improves mixing).  The kimwipe plug should enough to block solvent from passing back through the needle without pressure, but if you need to you can fully seal the leur lock/leur slip with an 8mm septa.  To effectively mix the beads most of the equipment in a chemistry lab won’t do (there are of course exceptions), so head over to the nearest biochemistry lab.  What you’re looking for is a rotating mixer, usually used to soak protein gels prior to visualization.  If the biochemists aren’t terribly keen on your syringes mixing with their equipment you can pick up one of your own from Fisher, with the cheapest around $500.  The stage is large enough to hold about eight syringes at once, and setting up that many reactions will keep you busy all day.

Once your beads have swelled, drain off the excess solvent [1].  From here the chemistry is the same as during large scale, just in a new container.  I usually premix my solutions in five dram flasks, at least double labeled with the coupling mix of choice; once on the vial, once on the cap.  Cleavage is effected with a 2 hour exposure to 95:2.5:2.5 TFA:H2O:TIS (so long as your peptide is Arg/Cys/Met/Trp free).  While the resin is cleaving replace the needles with rubber septa, as there’s a bit of acid sensitive glue holding the metal portion of the needle in place.

After cleavage the dissolved peptides are either drained into a round bottomed flask and concentrated (wash the beads with 95:5 TFA:water, water or pure methanol) or transferred to a 5 dram vial and precipitated with ether (wash the beads with 95:5 TFA:water only, and don’t try this with hydrophobic or amphiphilic peptides).

Next up: Purification on the cheap with the poor man’s HPLC column.

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[1] One word of caution.  Don’t fully drain the syringe while the peptide is still on the beads.  The beads can be crushed between the plunger and kimwipe plug.

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