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Sigman Group Welcome Kit

Version 2.1

Guidelines, Procedures, & Requirements

SIGMAN GROUP University of Utah

Updated 08/30/2015

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1. Laboratory Safety

1.1. General Considerations

1.1.1. You may not work in the Sigman lab until receiving basic safety training by the Safety Officer. They will review with you a checklist of key safety issues and you will need to sign a form confirming that you have a complete understanding (and will follow) these procedures.

1.1.2. Lab goggles/glasses MUST be worn at all times while working in the lab! This is extremely important because even things that seem pretty common and safe (e.g., using the rotovap) involve placing glassware under reduced pressure, which can lead to implosions. If you wear glasses, the department will pay for prescription safety glasses – contact Rich G. for details.

1.1.3. Lab coats are required whenever working with or around hazardous and/or potentially flammable substances in the laboratory.

1.1.4. Do not wear gloves or your lab coat at your desk or in the group room.

1.1.5. You must receive training before using any pyrrophoric materials (Grignard reagents, Li reagents, alkyl Zn reagents) in the laboratory. Check with the group Safety Officer to arrange the training.

1.1.6. Know where all eyewashes are located in each lab.

1.1.7. Know where all safety showers are located in each lab.

1.1.8. Know where all fire extinguishers are located in the lab, what kind they are, and what they can be used for.

1.1.9. Nothing should be stored on the lab floors! Keep the floors free of anything other than lab stools and the 10 L LN2 dewar.

1.1.10. Flammable liquids at all times should be stored in the fume hoods or the flammable cabinets only. (Not more than 500ml of organic waste should be accumulated at the bench working space or at the hoods)

1.1.11. Do not to work alone (especially late at night) in the lab (computer work is okay). It is particularly critical that others are around and are informed about what you are doing if you are conducting a large scale-up (>5x the scale that you have conducted the reaction previously), running reactions with very reactive materials (i.e., strong oxidants or reductants, Grignard reagents, lithium reagents, etc), carrying out reactions requiring high pressure, or when running a reaction for the very first time. However, note that even something as seemingly simple as using the rotovap can lead to hazardous situation (chemical spill, implosion, etc). Also it is critical to avoid quenching or dispensing large quantities of highly reactive chemicals when no one else is around.

1.1.12. Always clear your bench space and hoods of any used or unwanted chemicals and flammable liquids before leaving. Especially, a common bad habit is to run a column and leave the test tube rack filled with flammable liquids open overnight. DO NOT DO THIS!!!

1.1.13. The last person out of the lab should turn all lights out and lock all doors.

EMERGENCY PROCEDURE: For Fire, Police, Ambulance dial 911 on your mobile phone and 9911 on desk phones – this will connect you to the campus emergency center.

Give your name, location and phone number.

Describe exactly what happened and what you need (e.g., ambulance, fire trucks,

police, etc.)

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Otherwise, they will send the campus police over to find this out first, which takes lots of time.

Do not hang up until asked to do so.

Other emergency services:

University Police: 801-585-2677

University hospital ER: 801-581-2291

Poison Control: 1-800-222-1222

For more emergency response contacts look here:

http://ehs.utah.edu/emergency-preparedness/emergency-contacts

After performing the emergency procedure contact a more senior person of the lab and lab safety officer (Harsh-9897088876).

1.2. Reaction Safety

1.2.1. LABEL, LABEL, LABEL all your reactions clearly! This is not only for your safety, but for everyone else’s as well.

1.2.2. Reactions under high pressure (e.g., with condensed gases or in super-heated solvents) are explosion hazards and should be treated with extreme caution. A blast shield should be placed in front any system larger than an NMR tube under pressure and/or a 4 mL vial. NMR tube reactions should also be treated with extreme caution – the hood sash should always be lowered when NMR tubes are under pressure.

1.2.3. Water condenser hoses should be fastened with Cu wire, and water flow should be turned as low as possible at night (water pressure increases at night).

1.2.4. Although vacuum is used all the time in the lab (for filtration, running the rotovaps, etc), you should keep in mind that this involves reduced pressures and therefore pose a significant implosion hazard. Use caution when evacuating any flask [especially large round bottoms and large filter flasks (>500 mL)] and check glassware regularly for cracks.

1.2.5. Exercise caution in pulling tubing off Schlenk ware! If it is too difficult to remove the tubing, carefully cut the tubing away with a razor blade. Excessive jerking and pulling will snap the glass stopcock off, which may result in a cut to your hand. Remember rubber tubing is inexpensive, and is meant to be cut when necessary.

1.3. Common Explosion Hazards

1.3.1. Oxidants (e.g., bleach, CrVI

and MnVII

salts, hypervalent iodine reagents, H2O2, etc) should be placed in separate waste from organic reagents/solvents. The oxidation of organics with these reagents can lead to violent exotherms/explosions.

1.3.2. Oxidizing acids (e.g., nitric acid and aqua regia) can react extremely violently with organics (especially acetone), and the resulting explosions/release of corrosive solutions can lead to serious injury. Acids should always be stored in a separate location from organic chemicals. Additionally, waste bottles for acids should be clearly marked and placed in a separate location from organic waste. This will prevent mistakenly pouring acid waste in with organics (which is the most common cause of this type of explosion).

1.3.3. Perchlorate salts can explode without warning, especially when concentrated in the presence of organics (once again, ClO4

– is a strong oxidant!). Always use a blast shield when

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concentrating mixtures containing these salts and avoid the use of the ClO4– counter anion

whenever possible.

1.3.4. Peroxides: Peroxide forming chemicals are considered to have a limited shelf life for safe use. One should avoid using long stored ether solvents (for eg: dioxane). http://ehs.berkeley.edu/lesson-learned-peroxide-explosion-injures-campus-researcher

1.3.5. Azide containing molecules should be handled with utmost precaution. They are energy-rich molecules and can be also heat and shock sensitive. https://web.stanford.edu/dept/EHS/prod/researchlab/lab/safety_sheets/08-203.pdf

1.3.6. Metallic lithium should never be placed in N2 filled dry boxes or under a nitrogen atmosphere on your line. A violent and highly exothermic reaction will result from spontaneous “Li3N” formation.

1.3.7. Remember that something as common as flash chromatography columns are run under high pressure and can crack/explode unexpectedly.

1.3.8. The condensation of liquid O2, liquid N2 and solid Ar in traps on your vacuum line can lead to explosions. See the vacuum line safety section for further details.

1.3.9. Any new chemical to be handled should be cautiously noted for its properties such as boiling point (eg: acetaldehyde, ethylene oxide etc.) functional groups (eg: azides) , heat and shock sensitivity (for eg. Diazomethane: highly explosive due to its extreme shock sensitivity, not to be handled with needles or glass apparatus with cracks), high flammability (eg: ethers) and pyrophoric properties (see Page 18 for safe use and handling of pyrophoric reagents).

1.4. Toxicity Hazards

• Thallium salts (e.g., TlOEt).

• Alkyl mercury salts (e.g., HgMe2).

• Tin reagents (especially tetra-alkyl or tri-alkyl aryl Sn compounds).

• Alkylating agents (e.g., MeI).

1.4.1. Exercise extreme caution when using these reagents!! Clean up spills in your hood and in public areas (balances, dry boxes, etc) immediately using appropriate procedures, and dispose of cleaning supplies/gloves in solid waste containers beneath the hood (to avoid fume inhalation).

1.4.2. Dispose of gloves (in solid waste container) whenever you may have come in contact with these reagents.

1.4.3. If any of these compounds are used in the dry box, be sure to (i) use a secondary pair of gloves so as not to contaminate the main gloves, (ii) dispose of all contaminated waste in a separate Ziploc bag before removing it from the box, and (iii) purge the box after the use of these compounds (and before opening the antechamber).

1.4.4. For specific instructions on how to wash glassware that has contacted these reagents, speak with Matt and/or a trained group member directly.

2. Vacuum Line Procedure—Using the Vacuum Line

You will receive thorough training on vacuum line technique by Matt and/or a trained group member before using your line. However, remember that many of the techniques involved can be confusing, and the consequences of making a mistake can be very

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dangerous as well as costly. Therefore, if you are ever in doubt about how to do something, please be sure to ask before proceeding.

2.1. The following are references that contain a lot of useful information on almost all aspects for Schlenk and high vacuum technique (i) Experimental Organometallic Chemistry, Andrea L. Wayda and Marcetta Y. Darensbourg, Eds., American Chemical Society: Washington DC, 1987. (ii) The Manipulation of Air Sensitive Compounds, D. F. Shriver, Robert E. Krieger Publishing House: Malabar, FL, 1982.

2.2. Argon is a solid and N2 is a liquid at LN2 temperature (–195 ºC). Therefore, it is extremely dangerous

to place LN2 cooled flasks/traps under Ar or N2 as significant quantities of these gases can condense. The huge pressure increase as the condensed Ar/N2 warms and moves to gas phase can produce extremely violent explosions. As such, never backfill a flask a LN2-cooled flask with N2/Ar and/or leave it under a flow of N2/Ar.

2.3. O2 condenses as a bluish liquid at LN2 temperature (–195 ºC). Liquid O2 can condense in traps if the line is opened to air for any period of time and can spontaneously explode when co-condensed with organics. As such, ALWAYS evacuate traps before placing them in LN2 (to remove air) and ALWAYS remove LN2 before venting traps to air.

2.4. Exercise caution when evacuating any flask on the vacuum line. Large round bottoms and solvent bulbs (>500 mL) are especially significant implosion hazards and should be evacuated with the hood sash down. Also, regularly check glassware for cracks and remove and clearly label defective glassware. (However, the glassblower can often fix it, so don’t just throw it away).

2.5. Using solvent pots/flasks on vacuum line: Attach flask/solvent pot to line via tygon tubing. Be sure to use grease where appropriate. Evacuate for two-five minutes and refill with N2. Repeat this cycle three times. At this point all of the air should be out of your system and the flask/solvent pot can be opened to N2 flow. Note: If the system won’t pump down below 10 mbar, there is likely a leak, and all joints should be checked and re-greased if necessary. Do not simply proceed – the solvent pots are shared, and contamination will cause problems for everyone!

2.6. Remember that all reactions/flasks that are open to N2 can and do “see” and cross-contaminate each other. As such, only “compatible” reactions should be open to N2 simultaneously, and care should be taken to avoid this situation if possible. When in doubt, please talk with Matt before proceeding.

2.7. A critical point: When using the group solvent pots (taking solvents into the glovebox), ALWAYS close off other reactions from the N2 and thoroughly flush your Schlenk line with N2 before exposing the pot to the N2 atmosphere. If necessary, use the solvent pot on someone else’s line rather than risking contamination. Remember – these pots are shared, so we have to keep them clean for everyone!!!. *When using the group solvent systems avoid

Traps

2.7.1. The large trap (closest to the pump) should be cooled with LN2 any time you are removing more than 1 mL of solvent under vacuum.

2.7.2. If you are leaving something on the vacuum line overnight, be sure to fill the traps right before you leave and right when you arrive in the morning. Generally, the LN2 will only last ~12 hr.

2.7.3. If not being used, the traps should be taken down at the end of the day, and the remaining LN2 should be returned to the group 10 L dewar.

2.7.4. Before putting traps back up, be sure that they are completely free of solvent (if necessary place them in the oven for 15-30 minutes before proceeding).

2.8. Pumps and Pump Oil

2.8.1. Pump oil should be changed three times a year – in December, April, and August. It is your responsibility to keep your pump clean (by avoiding contamination with solvents) and to change your pump oil on a regular basis. Remember, a clean pump will work smoother, longer, and most importantly it will pump down faster.

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2.8.2. For problems with your pump (poor pump performance, leaking, strange noises, etc), immediately shut it down and talk to Matt in order to diagnose the problem.

2.8.3. Familiarize yourself with your pump by reading the operating manual. This will be extremely helpful when it comes time to change your pump oil.

3. Cleaning Glassware

Note: Although it may not seem that important, cleaning glassware is one of the most important tasks that you will do in lab – contaminated glassware (along with contaminated solvents) are the two biggest causes of reactions going bad!

3.1. General Group Glassware

3.1.1. Rinse out flask into organic waste to remove organic material by washing with a H2O-miscible organic solvent like acetone, MeOH, or THF, depending on solubility.

3.1.2. Thoroughly clean grease off of all joints with hexanes and a Kimwipe.

3.1.3. Scrub both the interior and exterior of the flask vigorously with a washing brush and soap/warm water to remove salts and remaining residues.

3.1.4. Glass and Teflon stopcocks should be removed from joints before cleaning. They are easily damaged by small particles such as salts and the stopcock bore tends to hold up liquids.

3.1.5. Rinse flask with warm water (at least 2-3 times) and with distilled water (at least 2-3 times) to remove all soap/residues.

3.1.6. Finally, rinse with a small amount of acetone and place on the drying racks.

3.1.7. If glassware remains visibly dirty after this procedure DO NOT leave it on the drying rack for someone else to take and use!! ASK Matt about the best way to get it clean – this will usually entail either placing it in the base bath and/or washing with strong acid (e.g. conc. H2SO4, HNO3) to remove residual metal salts.

3.2. Frits

3.2.1. Rinse your frit with solvents in which the solids on it are soluble. Typically this would involve MeOH followed by acetone then EtOAc then CH2Cl2. Then, turn the frit upside-down and rinse with these solvents a second time.

3.2.2. Note that aqueous washes (i.e., those with bleach/water/acid/etc) are sometimes necessary to remove toxic reagents like Sn and/or other reagents that are soluble in these media. However, these washings need to be separated from the organic washings, and disposed of separately (see waste section). Also, washes with H2O and/or aqueous solutions should be followed by copious rinsing with MeOH before the introduction of immiscible organics like CH2Cl2 or hexanes.

3.2.3 If residue remains (especially metal-based residue) it can often be removed by placing 50% conc. HCl and 50% MeOH in the frit and allowing it to drip through slowly, followed by rinses with HCl, H2O and MeOH.

3.2.4 If any particulate matter remains on the frit and/or it is not completely white, you should place it in one of the bucket in for cleaning with piranha solution (conc. H2SO4/H2O2) or aqua regia (HCl/H2SO4). However, YOU MUST COMPLETELY REMOVE ORGANIC SOLVENTS from the frit before subjecting it to piranha solution or aqua regia (highly oxidizing!). So, rinse the frits with MeOH followed by copious water before placing them in the bucket

3.3. NMR Tubes

3.3.1. Rinse the contents of your NMR tubes into organic (or aqueous) waste (depending of the contents of the tube).

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3.3.2. Rinse tubes at least one to two more times with a wash bottle into your waste before using the NMR tube cleaner. These steps are important to avoid excessive contamination of the NMR tube cleaner with everyone’s samples.

3.3.3. Note that you should never stick the tip of a wash bottle into an NMR tube to wash it out. This will inevitably lead to breaking the end off the tube. Instead, always hold the bottle several cm away from the end of the tube to spray the solvent in.

3.3.4. If solids/precipitated metals remain in the tube at this point, clean it out with some solvent (typically acetone) and a pipe cleaner.

3.3.5. Use the NMR tube washer to finish cleaning the tube. Typical solvent rinses might involve MeOH followed by acetone, then EtOAc then CH2Cl2.

3.3.6. Note again that aqueous washings (i.e., those with bleach/water/acid/etc) are sometimes necessary to remove toxic reagents like Sn and/or when the reagents used in NMR experiments are soluble in these media. However, these washings need to be separated from the organic washings, and disposed of appropriately (see waste section). Also, washes with H2O and/or aqueous solutions should be followed by copious rinsing with MeOH before the introduction of immiscible organics like CH2Cl2 or hexanes.

3.3.7. Place NMR tubes flat in the oven to dry. However do not leave them in the oven for more than ~6-8 hrs (after which they should be placed in a dessicator for storage). Leaving the NMR tubes in the oven for longer than this can lead to warping, which may cause problems with spinning, shimming and/or result in breakage in the NMR instruments.

3.4. Syringes/Needles

3.4.1. ALL syringes need to be cleaned directly after use! This prevents them seizing up or clogging (often irreversibly) with dried out residues. Additionally, these expensive pieces of glassware are in limited supply and are not replaced often.

3.4.2. Clean gas-tight syringes by rinsing them 2-3 times with 3-4 different solvents. Typically this would include MeOH, acetone, EtOAc, and CH2Cl2.

3.4.3. Gas tight syringes should be placed in the oven after cleaning without their plungers for 3-4 hrs. Longer times in the oven can lead to cracking and/or damage to the syringe. They should then be placed in a desiccator. Plungers should be wiped off and placed directly into a desiccator after cleaning. This prevents irreversible expansion/contraction of the plunger from repeated heating/cooling cycles.

3.4.4. Non-disposable needles should be rinsed thoroughly using the aspirator vacuum needle cleaner with appropriate solvents (typically MeOH followed by acetone then EtOAc then CH2Cl2).

3.4.5. Once again, note that aqueous washing of both gas tight syringes and needles (i.e., those with bleach/water/acid/etc) are sometimes necessary to remove toxic reagents like Sn and/or when the reagents used are soluble in these media. However, these washings need to be separated from the organic washings, and disposed of appropriately (see waste section). Additionally, washes with H2O and/or aqueous solutions should be followed by rinsing with copious MeOH before the introduction of immiscible organics like CH2Cl2 or hexanes.

4. Glove Box

4.1. When to use the glovebox

4.1.1. To weigh out reagents stored in the glovebox

4.1.2. To conduct reactions and/or workups with extremely air-sensitive compounds that cannot be

handled using standard Schlenk techniques

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4.2. How to bring equipment into the glovebox

4.2.1. SIGN-IN

4.2.2. Glassware/spatulas/solvent bulbs

4.2.2.1.1. Three 2-3 minute Evacuation and Refill cycles of the small antechamber

4.2.3. Solid reagents

4.2.3.1.1. Any solid should be rigorously dried under vacuum

4.2.3.1.2. The solid should be placed in dried glass container covered with a rubber-banded

kimwipe

4.2.3.1.3. Three 2-3 minute Evacuation and Refill cycles of the small antechamber

4.2.4. Liquids

4.2.4.1.1. Solvent bulbs prepared according to section 5 can be brought in with three 2-3

minute evacuation and refill cycles of the small antechamber

4.2.4.1.2. Unopened never-punctured sure seal bottles should be electrical taped and three

VERY SLOW evacuation refill cycles should be done

4.2.4.1.3. Any other liquids require consultation with the glovebox managers

4.2.5. General

4.2.5.1.1. Nothing should be brought into the box when the box has been exposed to solvent

or while the box is purging

4.2.5.1.2. Once you complete your cycles, the chamber should be backfilled with N2 and

turned to the off position

4.2.5.1.3. Once you have brought your equipment into the box the door should be

immediately sealed and the chamber should be left under positive pressure of N2

4.3. Using reagents in the box

4.3.1. Solids

4.3.1.1. After weighing out any solids please return the chemical to its proper storage location

4.3.1.2. Clean up the balance and glove box floor using the brush(es) and supplied masking tape

4.3.1.3. Seal your flask

4.3.1.4. If you have waste that is pyrophoric you may seal it in a provided plastic ziplock back and

then take it to your hood for proper disposal. Please consult with Matt or the glovebox

managers if you do not know how to properly dispose of such waste.

4.3.2. Liquids

4.3.2.1. Before opening a liquid you must turn off the circulation in the box

4.3.2.2. You may only dispense one bulk solvent at a time.

4.3.2.3. Once you finish using one solvent, you must purge the box for the required time. You may

not work in the box while purging.

4.3.2.4. If you require using more than one solvent at a time you should

4.3.2.4.1.1. Dispense solvent A from the bulk container to a personal vial and seal

4.3.2.4.1.2. Purge

4.3.2.4.1.3. Dispense Solvent B from the bulk container to a personal vial and seal

4.3.2.4.1.4. Purge

4.3.2.5. Repeat for each required solvent.

4.3.2.6. Now you may use your personal supply of solvents at the same time.

4.3.2.7. This process prevents cross-contamination of bulk solvents

4.3.3. General

4.3.3.1. You should NOT bring in a solvent bulb to add solvent to a reaction that does not need to

be run in the glovebox. These solvents can be added using standard Schlenk techniques.

4.3.3.2. You MUST NOT expose any bulk solvent to a solvent atmosphere!!

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4.3.3.3. Once you are finished in the box, any running reactions should be sealed with Teflon caps

and wrapped with electrical tape or capped with an unpunctured septum before purging

4.4. Removing equipment/reactions/reagents from the box

4.4.1. The small antechamber should now be moved back to the off position

4.4.2. Anything you wish to take out should be placed in the chamber and sealed

4.4.3. Material should be IMMEDIATELY removed from the chamber and the chamber should be

evacuated.

4.4.4. Sign out

4.4.5. Purge the box if you have used solvents and mark accordingly in the log book

4.4.5.1. Change the pressure levels to 12 –low and 13 high

4.4.5.2. Open the vent at the top of the box

4.4.5.3. Wait

4.4.5.4. Close the vent

4.4.5.5. Return levels to 4 and 6

4.4.5.6. Turn the circulation back on

4.4.6. Remember to stop your purge, ask your bench mate to remind you if you have a problem

remembering

4.5. Storing chemicals in the glovebox

4.5.1. Solids

4.5.1.1. If you have personal solids that need to be stored in the glovebox you must contact the

glovebox manager for a bin and ALL of your personal equipment and chemicals should be

stored in that bin labeled appropriately

4.5.1.2. If your chemical must be kept in the freezer you will be assigned a space in the freezer

4.5.2. Liquids

4.5.2.1. Personal liquids should be stored in Teflon capped vials and secured with electrical tape.

These vials will then be stored in a personal jar with a Teflon cap which can be acquired

from the glovebox managers.

4.5.2.2. Bulk, non-volatile, non-coordinating solvents may be stored in the glovebox in solvent bulbs

with flat bottoms capable of standing upright.

4.6. General

4.6.1. No chemicals/reactions/vials/etc should ever be left on the glovebox floor when you are not

actively working in the box

4.6.2. Clean up after yourself and remove all trash as soon as you are finished in the box

4.6.3. Stocking the box

4.6.3.1. Vials/pipets/caps/kimwipes/jars/bins/lids are all provided by the glovebox managers.

4.6.3.2. DO NOT restock the box yourself

4.6.3.3. If you notice something is getting low, feel free to remind the managers, they will be

appreciative

4.6.4. If you need to use the large antechamber consult with the glovebox managers.

4.6.5. If you get a hole in the gloves, IMMEDIATELY notify the glovebox managers

4.6.6. When in doubt, ask for help!

4.6.7. Solvents that should never be used in the box

4.6.7.1. Methanol, acids, Chlorinated solvents, water

4.6.8. Chemicals that should only be used in the box in small quantities with rigorous care

4.6.8.1. Methyl triflate, methyl iodide

4.7. Carelessness Kills (i.e., When in doubt ask!)

4.7.1. Balances are sensitive instruments. When they are not level, the weights they read are not

accurate. The floor of the glovebox is uneven. When you move the balance, it is likely to not be

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level. If you want the weights you record to have useful physical meaning, then you don’t want

this. Don’t move the balance!

4.7.2. Open solvent bottles are bad in any case. They are especially bad in a glovebox because it is

nominally a closed system. In other words, the atmosphere saturates in solvent vapors. This

then leads to the catalyst bed becoming saturated in solvent vapors, which leads to shorter bed

lifetimes. Also, solvent contamination becomes a problem (ether, for example, gets into

everything easily).

4.7.3. Needles are sharp. They are the single largest hazard to the gloves. If you want to use a

septum with a needle through it to dry something under vacuum, please don’t push the needle

all the way into the septum. When you do this, the needle sticks out past the end of the septum,

just begging for a glove puncture.

4.7.4. Make sure that the trap is iced up before using the rotovap pump. Make sure that the pump is

on before opening any of the vacuum valves. Make sure that the trap is iced up while the pump

is on. Don’t leave the pump on if you aren’t using it. If the trap isn’t cold, the trap contents all

end up in the pump oil. If the pump isn’t on, using the vacuum system will result in

contamination of the glove box atmosphere with outside air.

4.7.5. Label and cap your samples. Unlabelled samples disappear sometimes. Uncapped samples

are almost certain to end up either in the trashcan or scattered all over the box.

5. Dri-Solv System

5.1. Everyone must be trained by Chun Z or David S before using the Dri-solv System.

5.1.1. ALWAYS keep the Nitrogen on, even when not using the solvent system. 5.1.2. Do NOT dispense a large excess of solvent into the receiving flask. Unlike a still, the excess

solvent cannot easily be returned to the solvent reservoir.

5.1.3. If you want dry, deoxygenated solvent, then you should use a syringe. Dispensing solvent through the drain valve without introducing air is not a trivial process.

5.1.4. Try to avoid backwashing, this can lead to inadvertent contamination of the receiving flask. 5.1.5. When in doubt, ask for instructions.

5.2. Solvent System Instructions (Old System): 5.2.1. See Figure 1 for a diagram of the receiving flask. 5.2.2. Using a Syringe: 5.2.3. Dry and degas syringe. If you don’t know how to do this, then please ask someone for help

instead of trying to use the solvent system alone. 5.2.4. Open the two dispenser valves to dispense solvent into the receiving flask. Refer to Figure 2 to

see the proper valve positions. 5.2.5. Close the valves once you think you have sufficient solvent. It can be hard to judge sometimes,

but dispensing extra solvent is wasteful so it should be kept to a minimum. 5.2.6. Increase the N2 flow to several (at least 4) bubbles per second. 5.2.7. Open the syringe valve and proceed to syringe out your solvent. Avoid backwashing if at all

possible. 5.2.8. Close the drain valve immediately afterwards. 5.2.9. Decrease the N2 flow to one bubble per 1-2 seconds. 5.2.10. Draining into a flask: 5.2.11. Open the two dispenser valves to dispense solvent into the receiving flask. Refer to Figure 2 to

see the proper valve positions. 5.2.12. Close the valves once you think you have sufficient solvent. It can be hard to judge sometimes,

but dispensing extra solvent is wasteful so it should be kept to a minimum. 5.2.13. Increase the N2 flow to several (at least 4) bubbles per second. 5.2.14. Open the drain valve and collect solvent in your flask.

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5.2.15. Close the drain valve immediately afterwards. 5.2.16. Decrease the N2 flow to one bubble per 1-2 seconds

5. GC/GCMS, HPLC and SFC

5.1. Everyone must be trained by Christine N/ Christiane S before using the GC.

5.2. Everyone must be trained by Celine S/ John G before using the HPLC.

5.3. Everyone must be trained by Margaret H before using the SFC.

5.4. Reservations. The GC’s and HPLC can be reserved up to one week in advance by signing up on the list on the front of the instruments. During the days, the GC can be reserved for up to two hours at a time. Additionally, the GC’s can be reserved overnight for longer runs. During the day the GC can be used for longer than 2 hours if no one else is using the GC. If after 2 hours someone would like to use the GC, you must stop your run and let him/her use the GC.

5.5. Samples and Sample Preparation

5.5.1. All samples placed on a GC should have your initials on the vial, or if running a large number ( >20 vials) at least have your initials on the first and last vial in the sequence.

5.5.2. All samples should be removed after your run is complete.

5.5.3. If you find samples left on the GC by the previous user, the vials should be moved to the vial holder underneath the autosampler.

5.5.4. Occasionally you may need more GC vials. Before taking brand new vials, you must first take any used vials that no one has claimed. These used vials can be easily cleaned and work as good or better than new vials.

5.5.5. Samples for GC should be prepared in the following manner.

5.5.5.1. ~1mg/ml concentration

5.5.5.2. Reagents incompatible with the GC should be removed either by workup or running the sample through a short plug of silica. Metals, water, and any other reagents that can damage the columns should be removed.

5.5.6. Samples for HPLC should be prepared in the following manner.

5.5.6.1. ~1mg/ml concentration

5.5.6.2. Only sample which have been purified should ever be injected on the HPLC. Severe damage to the columns will result if metals etc… are injected. The columns cost ~$1500 each!

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5.5.6.3. The sample should be free of any particulate matter before injection otherwise the HPLC can be clogged.

5.5.7. Samples for SFC should be prepared in the following manner.

5.5.7.1. ~1mg/ml concentration

5.5.7.2. Only sample which have been purified should ever be injected on the HPLC. Severe damage to the columns will result if metals etc… are injected. The columns cost ~$1500 each!

5.5.7.3. Samples should be diluted in an alcohol or alcohol/hexane mixture. Other solvents are not tolerated by the columns even in small amounts.

5.5.7.4. No particulate matter in samples.

5.5.8. GC Care and Maintenance

5.5.8.1. Before your run, make sure the wash vials are full and the waste vials are empty.

5.5.8.2. Baking off the column before your run can often lead to better results.

5.5.8.3. If your compounds are very sensitive or if there is little separation between peaks, you may want to replace either the glass wool plug or the septa in the injector port.

5.5.8.4. Running a bake after your samples have finished is usually a good idea.

5.5.8.5. Purifying all samples (e.g. removing catalysts) as much as reasonable will increase the quality of your GC data.

5.5.8.6. Always run a shut down method.

5.5.9. HPLC Care and Maintenance

5.5.9.1. Solvent: When using the chiral columns a solvent ratio of 80:20 hexanes:isopropanol should not be exceeded. If necessary 25% IPA will be tolerated but be sure to equilibrate the column back to a higher hexanes proportion before stopping the pump.

5.5.9.2. Flow rate Flow rate should be at 1mL/min. or less. A flow of 1.5 mL/min. will be tolerated if necessary.

5.5.9.3. Do not let the columns run dry! Be careful to check the solvent height prior to use. If you would like to start a sequence prior to leaving the lab, a command is available to shut down the lamp, pump, and column chamber electronics. In the sequence table screen there is a box labeled post run macro, if you check this and then select the option ‘shutdown.mac’ from the dropdown menu the lamp, pump, and column components will shutdown after the last run is complete, but make sure your run is longer than it normally would be to include flushing of the column.

5.5.9.4. Flush the column for 20 minutes or so, more if necessary before stopping the pumps.

5.5.9.5. If after a run you do not observe the expected product peaks you must increase the polarity of the solvent and pump the column clean. Do not consciously leave samples on the columns!

5.5.9.6. If you encounter any irregularities or problems in operating the HPLC contact the group member who is overseeing the instrument. Don’t try to troubleshoot the problem without someone knowing what you are doing.

5.5.10. SFC Care and Maintenance

5.5.10.1. Before operating the machine make sure the lamp is on and the black knob at the back of the Fluid Delivery Module is pointing towards the ceiling.

5.5.10.2. You must prime the pumps before starting the machine. If you do not know what this means contact the group member overseeing the SFC

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5.5.10.3. Always check the solvent levels, if there is any question fill them up.

5.5.10.4. Use previously determined methods for the SFC, if you require a new method contact the person overseeing the instrument.

5.5.10.5. If you switch the CO2 tanks you must immediately contact the person overseeing the instrument so a new tank can be ordered

5.5.10.6. When running a sequence you must always select “Stop Sequence” at the end of sequence command prompt, otherwise the machine will keep running indefinitely.

5.5.10.7. When any problem occurs you must first contact the person in charge of the instrument before attempting any troubleshooting.

6. Lab Notebooks

6.1 Maintaining a clear, well organized, and up-to-date lab notebook is critical for (a) keeping track of your experiments for your thesis, (b) any publications/ patents that you will write and (c) enabling future generations of students to reproduce your work.

6.2 General instructions for keeping a lab notebook are as follows.

a. Skip 3-4 pages in the beginning for the Table of Contents and update the TOC regularly (monthly, at least).

b. Use only non-erasable ink in your notebook.

c. Write the reaction/experiment clearly at the top of each page. If you are following a published procedure, indicate the reference from which the procedure was obtained.

d. Make a table including each reagent, it’s molecular weight, the measured quantity – g (or mL), mol, and eq – used in the reaction, and the commercial source/purity of the reagent.

e. Write a detailed experimental, including the rate/order/time/temperature of addition of each reagent and solvent, and, where appropriate, any color changes that take place during the reaction. Also, detail all work up procedures and TLC data (where appropriate) for the reaction.

f. Be sure to weigh the product and determine the % yield for all reactions!!

g. NMR spectra should be saved and labeled according to the notebook number, page, and sample they refer to. For example, 1mss23.007 would refer to Matt S. Sigman notebook #1, p. 23, sample #7.

h. Everyone is responsible for backing up their data.

6.3 A sample lab notebook page is shown at the end of this document.

7. General

7.1. Please notify Matt if you will be out of town for one working day or more.

7.2. Weekly group meetings will be posted every semester on the web – please notify Matt if you cannot attend for any reason.

7.3. It is important to keep up on the current literature in organic and organometallic chemistry – particularly as it relates to your project. Additionally, you will periodically be asked to choose a paper from the current literature to present in group meeting. The following are journals that you should read each week and are appropriate sources for group meeting papers:

J. Am. Chem. Soc. Organometallics

Org. Lett. J. Org. Chem.

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Angew. Chem., Int. Ed. Nature Chemistry Chemical Science Chem. Commun.

Science Nature

*Note that reading the literature is critical not only to learn more about your project/area of research but also to get you prepared for upcoming seminar speakers, proposal writing, orals, local and national ACS meetings, writing your own papers, and ultimately getting a job!

7.4. General tips for reading the chemical literature:

7.4.1. You cannot expect to read everything.

7.4.2. Try to read papers that are (i) the most interesting to you and (ii) the most relevant to your and the group’s research projects.

7.4.3. No one has time to read the entire text of every article. Read the abstract and introduction and then try to discern the major point of the paper from the Figures and Schemes. If you find something especially interesting or unclear consult the text for further details. Keep in mind when writing your own papers that these are the sections that are usually the most read.

7.4.4. Whenever possible, discuss with others what you have read! This will solidify your general knowledge as well as improve your understanding of what you have read.

7.4.5. Take particular note of papers that describe selective reactions. These are the most useful in synthetic chemistry and the most difficult to find by traditional searching techniques.

7.4.6. Keep an eye out for molecules that could be assembled using the methodology that you are developing. This will be helpful for those of you who are interested in applying methodology in total synthesis, as well as for writing proposals.

7.5. Other journals to keep an eye on (monthly) are:

Tetrahedron Letters Tetrahedron

OBC Dalton Transactions

Chem. Reviews Acc. Chem. Res.

Synlett

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Sigman Group

Research Notebook Guidelines

Your notebook should be clear and complete enough to permit an expert outside person (e.g. another group member) to repeat any of your experiments or to locate a given set of analytical data in your absence.

Enter all experiments, even those that did not work the way you wanted them to. This is important to elucidate trends from even poor results.

Maintain an up-to-date table of contents.

Take time every night to update your notebook before you go home. It is not only the best way to record what you have completed while it is fresh in your mind, but it is also a good way to organize your thoughts for the next day.

Record ideas related to your research into your notebook. It is good practice to sign and date them.

If you are screening multiple reactions and use a spreadsheet to tabulate your data, paste in a copy of this sheet. Additionally, if you are measuring rates of a reaction, you should paste in a copy of the plotted or raw data. The notebook should include the name of the computer file where the raw data is saved.

All your notebooks and your analytic data should be easily found (on your desk or shelves). You should not take your notebook home with you.

Keep in mind that your notebook and your data are the only legal records of your research. They are also the legacy you will ultimately leave to the group.

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(1) Date: When you

begin the experiment.

(2) Title or Goal: A

brief statement

indicating what you are

trying to accomplish.

(3) The Reaction: Use

structures whenever

possible. Only indicate

the product if it is a

literature procedure or

a reaction you know is

successful.

(4) Reference:

Literature or notebook

reference indicating

what procedure you

followed.

(5) Reagent Table:

Show amount, # of

mmol, molecular

weights, # of

equivalents, and source

of reagents. All

reagents or solvents

that were subjected to

any manipulations prior

to reaction (distilled,

recrystallized etc..)

should be specifically

noted.

(6) Experimental:

Write clearly in a

narrative or a list of

actions taken.

Carefully note the

manner in which

reagents were

combined (e. g. “added

over 20 min via syringe

pump”, “cannulated

into the flask”, etc…).

Also, provide detail of

the workup of the

reaction.

(7) Continuation:

note where

the experiment is

continued

(1) (2) Formation of Oxazoline.

(3)

(4) Reference or previous page of experiment.

(5)

(6)

(7) next page

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(8)

Purification:

Include detailed

information

regarding

purification.

This may

include TLC

conditions,

distillation

temperature, or

recystallization

solvents and

conditions.

This is vital for

reproducibility.

(9)

Conclusions:

Note yield and

also note any

analytic data

about the

compound

(NMR taken,

mp, GC, etc…)

to confirm

purity. Also,

sign and date

your notebook

on completion

of your

experiment.

(8)

(9)

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8. Handling Liquid Pyrophoric Reagents:

Pyrophoric liquids are liquids that can ignite spontaneously upon contact with air or moisture. Because of their high reactivity, special precautions must be taken when using them. The most important safety consideration is to protect them from air and moisture. Only trained researchers should handle pyrophoric liquids; in addition to reading this document, you must be trained by an experienced researcher before you use pyrophoric reagents.

Examples of pyrophoric reagents:

alkyl lithium reagents (RLi)

Grignard reagents (RMgX)

alkyl zinc reagents (R2Zn, RZnX)

alkyl aluminum reagents (R3Al, R2AlX, RAlX2)

LiAlH4 and DIBAL-H solutions

organoboranes (R3B)

BH3

LDA (LiNiPr2)

phosphines

silanes Particularly hazardous pyrophoric liquids include:

tBuLi

sBuLi

10 M nBuLi

R3Al (especially when neat)

neat DIBAl-H

http://www.chemistry.ucla.edu/sites/default/files/safety/sop/SOP_Organolithium_Safety.pdf

Laboratory Requirements: Use of more than 5 ml (for liquids) or 1 gram (for solids) of any pyrophoric substance should be registered in the Pyrophoric Lab manual available at Harsh’s desk. Fill in the details of the reaction and the person who would be your assistant in case if there is any fire caused while or after handling the pyrophoric substance. Note that quenching of a pyrophoric reaction can be equally dangerous as handling the reagent, especially during a scale up. Thus, plan your reaction such that you will not be alone when it is time to quench the reaction.

Note: Learn to use a fire extinguisher before there is an actual need to use it. (Be Prepared!!!)

8.1 General:

1. First time handling a pyrophoric material (Grignard, alkyl lithium, alkyl zinc, metal hydrides) in Sigman lab:

a. You must have a partner with experience working with pyrophorics working with you during the entire procedure (from the beginning through quenching of reaction)

b. You must write up a full, detailed procedure for what you plan to do (including scale, detailed experimental for adding reagents, detailed experimental for queching the reaction before starting in the lab. This procedure will be reviewed and discussed with your partner and with Matt before you start the experiment.

c. You must clear your hood space of all flammable liquids around it. No squirt bottles or Organic waste should be present during the handling and quenching of the pyrophoric reagent. In case you are sharing a hood with someone, let the other person know that you are planning an experiment with a pyrophoric reagent and any flammables may be required to be removed from the hood for a certain period.

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Note: It is more often that the flammable liquids around the reaction which can catch fire are more responsible for causing serious damages and injuries rather than the pyrophoric reagent itself.

2. ANY TIME handling tert-butyl lithium (even if you are an experienced pyrophirics user):

a. You must have a partner with experience working with pyrophorics working with you during the entire procedure (start through quenching of reaction)

b. You must write up a full, detailed procedure for what you plan to do (including scale, detailed experimental for adding reagents, detailed experimental for queching the reaction before starting in the lab. This procedure will be reviewed and discussed with your partner and with Matt before you start the experiment.

c. You must clear your hood space of all flammable liquids around it. No squirt bottles or Organic waste should be present during the handling and quenching of the pyrophoric reagent. In case you are sharing a hood with someone, let the other person know that you are planning an experiment with a pyrophoric reagent and any flammables may be required to be removed from the hood for a certain period.

Always:

1. Wear gloves, safety glasses, and flame retardant lab coat

2. Write scheme of the reaction being done, especially the pyrophoric reagent, on hood

3. Have appropriate fire extinguisher next to the hood

4. Remove all solvent bottles (squirt bottles & waste included) and flammables from hood

5. Clamp bottles and flasks securely

6. Alert lab room members of reaction being performed (and of any incident)

7. Use a luer-lock syringe

8. Use 1 1-2 ft long needle

9. Use syringes that are at least twice the volume of the amount of material to be dispensed.

10. Avoid using a syringe for quantities >10 mL of a pyrophoric.

11. Ask if you are unsure or have questions about anything.

Strongly suggested:

1. Use calibrated addition funnel and/or graduated cylinder to cannula transfer reagent. This is required for volumes >10 mL and strongly suggested for smaller volumes as well.

2. Have container of dry ice/ LN2 nearby to quench needle or syringe if needed.

3. Have partner for reaction if wanted, feel unsure, or infrequent user. Never work alone (use the Buddy system)

Fire Extinguisher Tips:

Use yellow Class D (Metals) yellow extinguisher for solid metals such as Na, Mg, and metal hydrides. This

20

extinguisher can also be used for alkyl lithiums, but is not necessary for these reagents.

Class ABC red extinguisher can be used for alkyl lithium reagents and other solution based metals.

21

22

23

24

25

References Subchapter 7. General Industry Safety Orders Group 1. General Physical Conditions and Structures Orders Introduction §3203. Injury and Illness Prevention Program.

Group 16 Control of Hazardous Substances Article 109. Hazardous Substances and Processes §5191. Occupational Exposure to Hazardous Chemicals in Laboratories. §5194. Hazard Communication.

J.A. Schwindeman, C.J. Wolterman, R.J. Letchford: Chem. Health & Safety, May/June p. 6-11, 2002. See:http://membership.acs.org/c/chas /techarchive/organolithium%20paper.pdf

EXHIBIT:

Exhibit 1. Example of setup for extracting py rophoric chemical from reagent bottle.

Exact setup will depend on specific requirements. (Photo provided courtesy John Palmer of UC San Diego)

5

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Common TLC Stains:

Anisaldehyde: Slowly add 50 mL of concentrated sulfuric acid to one liter of 95% ethanol [best results when you cool the solution to 4 deg C and add the sulfuric acid portionwise over about 20-30 minutes]. Wait 5 minutes, then add 20 mL of commercial p-anisladegyde followed by 15 mL of glacial acetic acid.

Verghn's reagent: Dissolve 40 g of ammonium molybdate and 1.6 g of ceric sulfate in 800 mL of 10%

(v/v) sulfuric acid/ water.

Ninhydrin: Dissolve 200 mg of ninhydrin in 125 mL of 95% ethanol.

Phosphomolybdic Acid (PMA): Dissolve 25 g of phosphomolybdic acid crystals in one liter of 95% ethanol.

Potassium Permanganate Basic: Dissolve 3 g potassium permanganate and 20g potassium

carbonate in 300 mL water, then add 5 mL of 5% (w/v) of aqueous sodium hydroxide.

Vanillin: Slowly add 50 mL of concentrated sulfuric acid to one liter of 95% ethanol. Wait 10 minutes, then add 25 g of vanillin.

27

28

Characterization Data

Structure:

Notebook/Page #

Appearnce :

:

:

:

M. Pt./ B. Pt.

Specific rotation (solvent)

Concentration (g/ml)

Rf (Solvent)

Resolution :

:

IR: Low Res Mass Spec:

High Res Mass Spec:

Calculated: Found:

1HNMR: 13CNMR

Peak Mult. Coupling

constant

Assignment Peak Assignment