GUIDANCE MANUAL Model 900 BGI 900 HIGH VOLUME CASCADE IMPACTOR

GUIDANCE MANUAL

Model 900

BGI 900 HIGH VOLUME CASCADE IMPACTOR

BGI Incorporated 58 Guinan Street Waltham, MA 02451 USA Tel : 781 891-9380 Fax: 781 891-8151 www.bgiusa.com [email protected]

Version 1.0.0 December, 2008 Table of Contents

Patents, Copyrights and Trademarks ...... 1 Safety Notice ...... 2 Warranty (U.S) ...... 3 Operation manual Revision List ...... 4 Section 1. Introduction ...... 5 Section 2. Hardware Installation ...... 8 2.1 Description of the BGI900 ...... 8 2.2 Description of the Sampling Pump ...... 11 2.3 Description of the PUF Substrate & Filter Material ...... 13 2.4 System Setup ...... 15 2.4.1 Cleaning the Glassware & Tools ...... 16 2.4.1.1 Thermal Cleaning ...... 16 2.4.1.2 Solvent Cleaning ...... 17 2.4.2 Preparing the PUF Substrate ...... 18 2.4.2.1 Cleaning the PUF Substrates ...... 19 2.4.2.2 Storing the Clean, Dry PUF Substrate ...... 21 2.4.2.3 Cleaning the Substrate Holders ...... 22 2.4.2.4 Cleaning the Ultrafine Filter Hplders ...... 24 2.4.2.5 Preparing the PM-10 Grease Collection Platform ...... 25 2.4.3 Assembling the Collection Stages ...... 25 2.4.4 Installing the Sampler Onto the Tripod ...... 34 2.4.5 Setting Up the Pump ...... 36 Section 3. Post Sampling Analysis ...... 38 3.1 Removal of the Collection Platform ...... 38 3.2 Substrate Extraction for Analysis of Water Soluble Compmnents .... 42 3.2.1 Labware Washing ...... 43 3.2.2 Extracting PUF Substrates with Milli-Q Water ...... 44 3.2.2.1 PM-10 PUF Substrate (P1 Size) ...... 44 3.2.2.2 PM-2.5 PUF Substrates (P2 Size) ...... 46 3.2.2.3 PM-1.0, PM-0.5 & PM-0.1 PUF Substrates (P3 Size) ...... 48 3.2.2.4 Ultrafine Filter (Ultrafilter) ...... 50 3.3 Substrate Extraction for Toxicological Testing ...... 51 3.3.1 Labware Washing ...... 52 3.3.2 Extracting PUF Substrates with Methanol ...... 53 3.3.2.1 PM-10 PUF Substrate (P1 Size) ...... 53 3.3.2.2 PM-2.5 PUF Substrates (P2 Size) ...... 55 3.3.2.3 PM-1.0, PM-0.5 & PM-0.1 PUF Substrates (P3 Size) ...... 57 3.3.3 Evaporation of Solvent from PUF Extracts ...... 59 3.3.4 Extracting Ultrafilters with Methanol ...... 61 3.3.5 Evaporation of Solvent from Ultrafine Extracts ...... 63 3.4 Analysis of Samples ...... 67 References ...... 68

Patents, Copyrights and Trademarks

This instrument from BGI Incorporated (BGI) is covered by one or more of the following US patents: U.S. Patent Office Number 6,435,043 Harvard School of Public Health

This instrument is fabricated, distributed and technically supported by BGI Incorporated. The technical design is from the Harvard School of Public Health (HSPH) and BGI is the sole licensed manufacturer of the High Volume Cascade Impactor, BGI Model 900.

This Guidance Manual may contain trade secrets and confidential information proprietary to BGI and HSPH. The documentation and information contained therein may not be used, duplicated or disclosed to anyone, in whole or in part, other than as authorized by Agreement and with the express written permission of BGI.

Note: The former product name of this instrumentation was “CHEMVOL©” since 2003, and was formerly a registered trademark of Rupprecht & Patashnick Company Incorporated©.

Safety Notice

Repair of instrumentation manufactured by BGI Incorporated (BGI) should only be attempted by properly trained service personnel. Do not tamper with this hardware. Use established safety precautions when working with this instrument.

BGI and its licensed distributors can not foresee all possible modes of operation in which the user may attempt to utilize this instrument. The user assumes all liability associated with the use of the instrument. BGI and its distributors further disclaim any responsibility for consequential damages. Use of this product in any manner not intended by BGI, the manufacturer, will void the safety protection provided by the equipment, and may damage the equipment and subject the user to injury.

Warranty (U.S.)

BGI warrants equipment of its manufacture and bearing its nameplate to be free from defects in workmanship and material. We make no warranty, express or implied, except as set forth herein. BGI's liability under this warranty extends for a period of one (1) year from the date of BGI's shipment. It is expressly limited to repairing or replacing at the factory during this period and at BGI's option, any device or part which shall within one year of delivery to the original purchaser, be returned to the factory, transportation prepaid and which on examination shall in fact be proved defective.

BGI assumes no liability for consequential damages of any kind. The purchaser, by acceptance of this equipment, shall assume all liability for consequences of its misuse by the purchaser, his employees or others. No information derived from the use of this equipment is warranted for any purpose whatsoever. This warranty will be void if the equipment is not handled, installed, or operated in accordance with our instructions. If damage occurs during transportation to the purchaser, BGI must be notified immediately upon arrival of the equipment. The Equipment will be returned via collect shipment.

A defective part in the meaning of this warranty shall not, when such part is capable of being repaired or replaced, constitute a reason for considering the complete equipment defective. Acknowledgment and approval must be received from BGI prior to returning parts or equipment for credit. BGI makes engineering changes and improvements from time to time on instruments of its manufacture. We are under no obligation to retrofit these improvements and/or changes into instruments which have already been purchased.

No representative of ours has the authority to change or modify this warranty in any respect.

Operation Manual Revision List

As BGI instrumentation fabrication and design changes, so do our Guidance Documents and Operator Manuals. However, these changes may affect only one aspect of an instrument, while leaving the instrument as a whole unchanged. To explain these individual changes to BGI users, BGI will update only these sections of its Guidance Document or Operator Manual that are affected by the instrument updates or improvements. When this manual changes, so does it revision number. The Guidance Document Revision Number is located on the front cover.

The latest versions of any BGI manual may be found on our website: http://www.bgiusa.com/cau/manuals_2005.htm

To assist users of the High Volume Cascade Impactor track the future changes, the following list of manual sections with its respective revision number are listed:

Section Number Description of Change Date

Guidance Document “Chemvol© Model 2400 May 2003 (Discontinued)

(Replaced With) Model 900 High Volume Cascade Impactor December 2008

Section 1. INTRODUCTION

The High Volume Cascade Impactor (Figure1-1), manufactured by BGI Incorporated (BGI) is a licensed and patented instrument design of the Harvard University School of Public Health (HSPH). The initial design, testing and verification of the High Volume Cascade Impactor was conducted by HSPH. BGI is the sole licensed manufacturer and distributor of this product.

The BGI High Volume Cascade Impactor (HVCI) is designed to aerodynamically classify the particles in the ambient air by the principle of Cascade Impaction. A set of Orifice Stages with decreasing orifice diameters are placed sequentially on top of one another and ambient air is drawn through the instrument. The aerodynamic diameter separation is made at the exit of each Orifice Stage. Separation and collection of the following particles are possible at a sample flow rate of 900 Liters a minute:

Top Collection Stage PM10 Second Collection Stage PM2.5 Third Collection Stage PM1.0 Fourth Collection Stage PM0.5 Fifth Collection Stage PM0.17 Bottom Stage Absolute Filter

Figure 1-1. High Volume Cascade Impactor System with Mounting Tripod

The amount of particulate mass that must be collected on each stage for certain types of toxicological and chemical speciation sampling ranges from a milligram (mg) to a gram (g). To collect the amount of mass needed over a short period of sample event, the High Volume Cascade Impactor is operated at a designed high sample flow rate of 900.0 Liters/Minute.

Historical filter and impaction methods were not suitable for classifying particulate matter at high flow rates. For example, filtration methods such as the Hi Volume Sampler (Hi-Vol), which operates at a flow rate of 1,000 Liters/Minute, require large glass or quartz fiber filters (8x10inches, 20.3x25.4cm). As a result, these collection media also require large quantities of solvents to recover the collected particulate matter and the weighing analysis is conducted on an analytical balance rather than a microbalance. This severely limits the Hi-Vol usefulness for both toxicological and chemical speciation characterization studies.

In contrast, conventional inertial impactor designs, like the HVCI have the ability to focus the collected particulate matter onto relatively small surfaces. These small collection surfaces allow the operator to recover particulate matter from the impaction substrate surface using relatively small extract volumes of solvents.

As with all Cascade Impactor designs, the fast-moving particulate matter impacts a hard collection substrate and often some particles “bounce” and reenter the air stream and are carried off to the next or final filter collection stage. This is commonly referenced as “particle bounce” and is a limitation of most commercial cascade impactor designs, which causes errors in the collection design of the instrument.

To overcome these particle bounce problems, some impactors require the operator to coat the impaction collection substrate with a sticky substance, such as oil or silicone grease. However, the use of impactor oil and greased substrates limits the analytical ability of the instrument. The grease may not allow for chemical analysis of the particulate matter, and may possibly change the particulate matter characteristics of those particles that do not stick to the intended collection stage. In addition, for relatively large amounts of collected material, the substrate collection efficiency depends on the amount of particulate matter mass collected. Consequently, only relatively small amounts of particulate matter mass can be accurately accumulated on grease or oiled impactor substrates.

To overcome the particle bounce and low capacity issues of typical commercial cascade impactors, the Harvard School of Public Health designed the Model 900, High Volume Cascade Impactor with a unique Polyurethane Foam (PUF) impaction substrate.

The large pores and relatively low overall density characteristic of PUF allows particles to be collected using the conventional impaction process. Polyurethane foams produce impaction processes with negligible particulate matter bounce-off losses because particulate matter can impinge onto the substrate with a possible gradual decrease of particle velocity. Because of their highly specific surface area, these substrates present a high-collection capacity and can be used to collect milligram to gram quantities of particulate matter materials.

The High Volume Cascade Impactor can be used to collect ambient particulate matter samples for one week or longer (depending on how much particulate matter pollution exists in the ambient atmosphere) at a design flow rate of 900 l/min, (31.8 CFM). The impactor cut-points range from 0.17 to 10 µm. Smaller (ultrafine) particulate matter can be collected on a disk of absolute filter material located downstream of the final impactor stage.

Figure 1-2. Rain cap, four impactor stages and Polyurethane (PUF) foam collection substrates.

Section 2: Hardware Installation

This section describes the installation of the Model 900 High Volume Cascade Impactor. This section also covers a number of operational considerations.

2.1. DESCRIPTION OF THE MODEL 900 HIGH VOLUME CASCADE IMPACTOR

Figure 2-1 displays a schematic diagram of the sampler. This figure shows a configuration of all possible impactor stages followed by an ultrafilter. The key feature of the High Volume Cascade Impactor (HVCI) sampler is the ability to collect a large quantity of particulate matter and to classify the material according to particle size. The available selection of stages have cut points of 10 µm (PUF and grease), 2.5 µm (PUF), 1 µm (PUF), 0.5 µm (PUF), 0.17 µm (PUF). The sampler also has an optional final-stage ultrafine filter (“ultrafilter”) that collects ultrafine particulate matter that measure less than 0.1 micrometers. Figure 2-1. Schematic drawing of an installation of the Model 900 sampler.

TOP COLLECTION STAGE PM-10

PM-2.5 PM-1.0 INTERMEDIATE COLLECTION PM-0.5 STAGES PM-0.1 EFFLUENT BOTTOM FILTER AIR COLLECTION STAGES

BYPASS TRIPOD AIR

FILTER

PUMP

Figure 2-2 lists the actual D50 cut points for each of the HVCI stages. The ultrafine PTFE coated glass fiber filter stage has > 99% collection efficiency for particulate matter below 0.1 micron.

Figure 2-2. Actual D50 cutpoints for each of the Stage Actual Measured D 50 (µm) HVCI collection stages. PM-1 0 PUF 9.5

PM-1 0 Grease 9.9

PM-2.5 2.5

PM-1.0 1.0

PM-0.5 0.6

PM-0.1 0.17

The HVCI sampler’s design features a circular slit (sectioned into 3 equal arcs) acceleration platform, with corresponding PUF rings for impaction substrates (Figure 2- 3). The acceleration platforms are mounted in modular cylindrical housings called “collection stages” (Figure 2-3). The collection stages are stacked in sequence, with the selected stages in proper order (by descending cut-point sizes) (Figure 2-1). A removable rain cover is attached to the top stage (Figure 2-4), which always is PM- 10 grease or PM- 10 PUF substrate.

Figure 2-3. PM-2.5 collection stage and PM- 2.5 PUF inside a substrate holder (collection platform assembly). Note: HVCI is inverted. Acceleration platform

PM-2.5 PUF Collection stage

Substrate holder

Figure 2-4. HVCI Sampler.

The design of the collection stages includes:

• A top collection stage for the PM-10 cut point that has the appropriate fittings for attaching the rain cover to the system. This stage must contain the PM-10 (PUF or Silicone Grease) stage.

• Any number of subsequent intermediate stages to characterize the particulate matter in ambient air. These following stages may include PM- 2.5, 1.0, 0.5, 0.17, and ultrafine (“ultrafilter”).

• A final, bottom collection stage which can be PM- 2.5, 1.0, 0.5, 0.17, or ultrafine.

2.2. DESCRIPTION OF THE SAMPLING PUMP (Not Supplied by BGI)

The sampling pump recommended by BGI (Figure 2-5) is a Regenerative Blower, positive-displacement type pump. The North American vendor is GAST Manufacturing Inc. http://www.gastmfg.com/pdf/newblower/R4H.pdf See the pages labeled #46 and #47, Regenair® Regenerative Blower R4H Series, Model # R4H3060A-1 (Vacuum Only). There have been several safety features built into the pump to ensure high- quality results. Once the pump is run for a period of several hours, it should produce a steady operating rate that needs little adjustment.

Figure 2-5. HVCI pump used by the former ChemVol Model, Replaced with the Gast R4H3060A-1. .

The pump for the HVCI system is designed to accommodate a variety of different HVCI sampler configurations. The pump will pull as much air as mechanically possible. If the pump draws in excess of the specified 900 l/min, a bypass valve must be mounted on the pump inlet to increase the pressure drop (Figure 2-6), to maintain the flow at the desired rate of 900 l/min. The pump should give the user a free air flow amount capacity above the demand of the system.

Figure 2-6. A Typical bypass valve on the pump that allows adjustment of flow rate to the pump.

When you select a pump, you must choose a system that produces a minimum of 900 Liters/Minute with the final filter in place. The TX40 PTFE coated filter media alone produces a differential pressure of 24 inches of water (1.8inches of Mercury) at 900 l/min. When choosing a new system, be sure to specify sufficient extra blower pressure drop for contingencies. Two different blowers that are recommended by BGI that produce 50 and 150 inches H2O of column vacuum at 900 l/min. The Gast Pump is recommended for 240 Volt AC North American operation. The Ritchle Thomas Model SAH95 is recommended for 240Volt use in Europe. The desired flow through the system is 900 l/min ± 5%. Refer to Section 2.4.5 for further information regarding the pump setup and operation. Additional hardware will be necessary to make a pump assembly. These include a 1” heavy wall vacuum hose with connectors to fit the Model 900 1” male pipe thread and the 2” female Gast pump. The pump manufacturer or industrial supplier should be able to outfit the pump assembly, including vacuum hose, check filter, bleed valve, exhaust muffler and electrical switches and breakers to meet local codes.

2.3. DESCRIPTION OF THE PUF SUBSTRATES AND FILTER MATERIAL

The HVCI sampler uses polyurethane foam (PUF) as the impaction substrate (Figure 2-7). The PUF substrates use ether-based foam, which is more chemical- and temperature-resistant than other types of foam. The substrates come in three different sizes for the PM- 10, PM-2.5, and PM-1 to 0.1 stages. PUF volume and dimensions are explained in Figure 2-8.

Figure 2-7. PM-2.5 PUF substrate.

Figure 2-8. PUF volume Stage Siz and dimensions. Name Normalized Volume and Inside Diameter Outside Diameter Surface Area

PM-1 0 PUF1 3.5 inches 6.5 inches 6

PM-2.5 PUF2 4.5 inches 5.5 inches 2

PM- 1.0 PM-0.5 PUF3 4.75 inches 5.25 inches 1 PM-0. 1

The ultrafine stage (ultrafilter) (Figure 2-9) uses a PTFE coated Glass Micro-Fiber filter that traps particulate matter, essentially an absolute filter. These substrates and filters are available through BGI and are packaged in quantities of 10 (Figure 2-10).

Figure 2-9. Inverted Ultrafilter installed inside a collection stage with a spare ultrafilter .

Figure 2-10. PUF Name BGI Part Number substrate and ultrafilter part numbers. P1 HVCI7952 (PM-1 0)

Grease HVCI7888 (1 Gallon. can) P2 HVCI7953 (PM-2.5)

P3 HVCI7954 (PM-1.0 to PM-0.1)

Ultrafilter HVCI7955 (<0.1 µm)

2.4. SYSTEM SETUP

When the sampler is assembled and the clamps are securely tightened, there are no moving parts to manipulate. With an actual experiment in a specific location, you can gradually extend the sampling period from one day to multiple days. Be sure to closely monitor the PUF substrate(s) to prevent too much particulate matter from collecting on the substrate(s) and ultrafilter. If you allow too much particulate matter to collect on the substrate(s) and ultrafilter, it may change the cut point of the stage or cause bounce-off losses (Section 1).

To set up the HVCI sampler, you must determine the number of stages that you want to use, clean the glassware and other instruments used to prepare the PUF substrates (Section 2.4.1), prepare the PUF substrates (Section 2.4.2), assemble the collection stages (Section 2.4.3), install the entire system onto the tripod (Section 2.4.4), and turn on the pump and adjust its bypass valve (Section 2.4.5).

2.4.1. CLEANING THE GLASSWARE AND TOOLS

There are two methods of cleaning the glassware and other instruments that are used for loading the HVCI sampler and performing the subsequent analysis on the PUF substrates and ultrafilter. The two methods are thermal cleaning and solvent cleaning. If you are using Teflon labware, do not use the thermal cleaning technique. The thermal cleaning technique uses high temperatures that will damage Teflon labware and release toxic fumes, such as carbonyl fluoride.

2.4.1.1. THERMAL CLEANING

You will need two ovens for this procedure: a conventional oven that can reach temperatures of up to 100° C, and a high-temperature oven that can reach temperatures of up to 600° C.

Follow these steps to perform a thermal cleaning procedure:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Wash all glassware and other necessary tools with detergent. 2) Rinse the glassware and tools with distilled water. 3) Rinse the glassware and tools three times with Milli-Q water. 4) Place the glassware and tools in a conventional oven at 100° C until they are dry. 5) After the glassware and tools are dry, remove from them from the oven and wrap them with aluminum foil. 6) Place the glassware and tools that are wrapped in aluminum foil into the high-temperature oven. 7) Set the oven temperature to 450° C. 8) Turn on the high-temperature oven. 9) Bake the glassware and tools in the high-temperature oven for four hours. 10) Turn off the oven but DO NOT OPEN THE OVEN DOOR. 11) Allow the temperature of the high-temperature oven to drop to ambient temperature. 12) Open the oven door and remove the glassware and tools.

2.4.1.2. SOLVENT CLEANING

Follow these steps to perform a solvent cleaning procedure:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Wash all glassware and other necessary tools with detergent. 2) Rinse the glassware and tools with distilled water. 3) Rinse the glassware and tools three times with Milli-Q water. 4) Place the glassware and tools in a conventional oven at 100° C until they are dry. 5) After the glassware and tools are dry, remove from them from the oven. 6) Place the glassware and tools into a 4000 ml beaker. 7) Add Milli-Q water into the beaker until the water covers the glassware and tools. 8) Place the beaker into an ultrasonic bath. 9) Place the ultrasonic bath into a positive-flow clean-air hood. 10) Sonicate for 1 hour. 11) Repeat steps 6-10 using hexane instead of Milli-Q water. 12) Repeat steps 6-10 using methanol instead of hexane. 13) Repeat steps 6-10 using dichloromethane instead of methanol. 14) After you have sonicated the glassware and tools using dichloromethane, place the glassware and tools into a stainless steel basket. 15) Place the stainless steel basket into the clean air hood until the glassware and tools are dry. 16) After the glassware and tools are dry, wrap them with clean aluminum foil to protect them from dust and other contamination during storage.

2.4.2. PREPARING THE PUF SUBSTRATES

Prepare the PUF substrates and substrate holders in a laboratory. Be sure to fill the PM-10 grease platform until the grease is level with the top of the rim of the substrate holder. The PUF platforms only need to have the foam placed into the groove of the substrate holders. Be sure to line up the holes in the PUF substrates with the holes in the substrate holders. The PM- 10, PM-2.5, PM-1, PM-0.5 and PM-0. 1 PUF substrates and their substrate holders (collection platforms) will fit onto the standoffs located on the acceleration platforms.

2.4.2.1. CLEANING THE PUF SUBSTRATES

You must clean the PUF substrates before sampling to remove any trace quantities of materials that may have collected on the foam during the fabrication process. The cleaning procedure uses a series of four different solvents (purified water, methanol (CH3OH), dichloromethane (CH2Cl2), and hexane (C6H14) to ensure that the extraction process removes materials that are highly polar, slightly polar, and completely nonpolar. Be sure to use high-grade solvents for the cleaning procedure, and exercise great care while handling the solvents. Use a fume hood to minimize personal exposure to the solvents.

Be sure to sonicate only in glassware, because plastic tends to damp the ultrasonic energy too much. Organic solvents will cause the PUF substrates to swell. This swelling process will reverse as the PUF substrates dry. When the PUF substrates are in a swollen condition, they are more fragile than when they are dry.

Tools and materials required for this procedure: 1000 ml and 4000 ml beakers Un-serrated stainless steel forceps Ultrasonic bath that is large enough to hold the 4000 ml beaker Clean-air positive-flow hood (with filters to remove particulate and gaseous contaminants) Ultra-pure water (Milli-Q water or equivalent) Methanol Dichloromethane Hexane Clean stainless-steel basket (12.7 cm x 10 cm x 10 cm [L x W x H])

Follow these steps to clean the PUF substrates:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Place the P2 and P3 (Figures 2-8 and 2-9) PUF substrates into the 1000 ml glass beaker. Place the P1 PUF substrate into the 4000 ml beaker. 2) Add the Milli-Q water to the beaker until it covers the PUF substrates. 3) Place the beaker in the ultrasonic bath. 4) Place the ultrasonic bath in the clean air hood. 5) Son icate for 1 hour. 6) Repeat steps 2-5 using hexane instead of Milli-Q water.

7) Repeat steps 2-5 using methanol instead of hexane. 8) Repeat steps 2-5 using dichloromethane instead of methanol. 9) After you have sonicated the PUF substrates using dichloromethane, place the PUF substrates into a stainless steel basket. 10) Place the stainless steel basket into the clean air hood.

11) Leave the stainless steel basket in the clean air hood for 24 hours, or until the PUF substrates are dry. 12) After the PUF substrates are dry, go to Section 2.4.2.2 (Storing the Clean, Dry PUF Substrates).

2.4.2.2. STORING THE CLEAN, DRY PUF SUBSTRATES

The most important aspect of storing the PUF substrates is to prevent ambient contamination before the foam is used. Protection from particulate matter contamination is straightforward, but gas-phase contamination requires a container that prevents diffusion of contaminants. Foil bags prevent both particulate-matter and gas- phase contamination. Also, the foil bags keep the PUF substrates flat for use. Be sure to keep the PUF substrates from distorting during the storage process. Estimated shelf life for cleaned PUF substrates is six months.

Tools and materials required for this procedure: 1000 ml and 4000 ml beakers Unserrated stainless-steel forceps Aluminum foil For P2 and P3 substrates (Figure 2-10): 500 ml tall glass jars with PTFE Teflon-lined caps For P3 pieces (Figure 2-10): 4000 ml tall glass jars with PTFE Teflon-lined caps

Follow these steps to store the PUF substrates:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Clean and dry the PUF substrates (Section 2.4.2.1). 2) While the PUF substrates are still inside the clean-air hood, transfer the dry PUF substrates to the glass jar(s). 3) Tightly cap each jar. 4) Wrap each jar with aluminum foil (this is necessary to keep out light). 5) Store the wrapped jars in a refrigerator at 4° C.

2.4.2.3. CLEANING THE SUBSTRATE HOLDERS

Follow these steps to clean the substrate holders:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Locate the empty substrate holder(s) (Figure 2-11).

Figure 2-11. Empty substrate holder.

2) If you are cleaning new, unused substrate holders, go to step 3. If you are cleaning previously used substrate holders, go to step 6. If you are cleaning a previously used PM-10 grease substrate holder, go to step 7. 3) Clean all surfaces of the new, unused substrate holders with detergent, water and a brush. Go to step 4. 4) Rinse all surfaces of the new, unused substrate holders with distilled water. Go to step 5. 5) Dry all surfaces of the new, unused substrate holders with Kimwipes, or other other laboratory tissues. Go to step 10. 6) Clean all surfaces (that come in contact with the PUF substrates) of the previously used substrate holders using cotton swabs moistened with Milli-Q water. Go to step 10. 7) Remove the used grease with a Kimwipe. Go to step 8. 8) Wash the PM-10 grease substrate holder with detergent and water. Go to step 9. 9) Dry all surfaces of the PM-10 grease substrate holder with Kimwipes, or other other laboratory tissues. Go to step 10. 10) Store the substrate holders in plastic bags to keep them free from dust or other contamination. 22

2.4.2.4. CLEANING THE ULTRAFINE FILTER HOLDERS

Follow these steps to clean the ultrafine filter holders:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Locate the circular plate w/ bar and the screen (Figures 2-12, 2-13 and 2- 14).

Figure 2-12. Inverted Ultrafine collection stage with bar highlighted.

Figure 2-13. Ultrafine collection stage with retaining plate highlighted.

Figure 2-14. Ultrafine collection stage with Ultrafilter highlighted.

2) If you are cleaning a new, unused circular plate w/ bar and screen, go to step 3. If you are cleaning a previously used bar, circular plate and screen, go to step 6. 3) Clean all surfaces of the new, unused circular plate w/ bar and screen with detergent, water and a brush. Go to step 4. 4) Rinse all surfaces of the new, unused circular plate w/ bar and screen with distilled water. Go to step 5. 5) Dry all surfaces of the new, unused circular plate w/ bar and screen with Kimwipes, or other other laboratory tissues. Go to step 7. 6) Clean all surfaces (that come in contact with the ultrafine filter) of the previously used circular plate w/ bar and screen using cotton swabs moistened with Milli-Q water. Go to step 7. 7) Store the circular plate w/ bar and screen in a plastic bag to keep them free from dust or other contamination.

2.4.2.5. PREPARING THE PM-10 GREASE COLLECTION PLATFORM

Follow these steps to prepare the PM-10 grease collection platform:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Clean the PM-10 grease substrate holder (Section 2.4.2.3). 2) Fill the clean substrate holder with fresh grease (Dow Corning silicone high-vacuum grease). 3) Ensure that the entire volume (except for 0.25 cm at each end of the cavity) is filled with grease. 4) Using a metal straightedge, ensure that the height of grease is level with the height of substrate holder. When you fill the substrate holder with grease, it becomes the “PM-10 grease collection platform.” 5) Place the PM-1 0 grease collection platform into a glass container (crystallizing dish without a cover, 170 mm [ID] x 90-mm [H]). 6) Wrap the glass container with aluminum foil.

2.4.3. ASSEMBLING THE COLLECTION STAGES

After you have cleaned the PUF substrates and the substrate holders (Section 2.3.2), you must install the PUF substrates into the substrate holders. When you install a PUF substrate into a substrate holder, or install grease into the PM- 10 grease substrate holder (Section 2.4.2.5), the combined assembly becomes a “collection platform.” After you have prepared the collection platforms, you must install them into the collection stages. When the collection stages are assembled, you must then assemble the sampler (Section 2.4.3.2).

Follow these steps to assemble the collection stages:

NOTE: It is important to wear powder-free gloves while assembling the collection platforms. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Clean the PUF substrates and substrate holders (Section 2.4.2). 2) If you are using grease in the PM-10 collection stage, go to step 3. If you are not using grease in the PM-10 collection stage, go to step 5. 3) Clean the PM-10 grease substrate holder (Section 2.4.2.3). 4) Prepare the PM-10 grease collection platform (Section 2.4.2.5). Go to step 5. 5) Install the precleaned PUF substrates into the substrate holders to prepare the collection platforms (Figure 2-15). You may need to use the unserrated stainless steel forceps when installing the smaller PUF substrates into their substrate holders. The PM-1, PM-0.5 and PM-0.1 PUF substrates all use the same size foam ring (0.25 inches (6 mm) wide).

Figure 2-15. PM-2.5 PUF collection platform.

6) Ensure that the holes in the PUF substrate line up with the holes in the substrate holder (Figure 2-16). If you will be using the ultrafine filter (ultrafilter), go to step 7. If you will not be using the ultrafine filter (ultrafilter), go to step 12.

Figure 2-16. Lining up the holes in the PUF substrate with the holes in the substrate holder.

7) Remove the (4) four screws from the circular plate and remove plate (Figure 2-17). Go to step 8.

Figure 2-17. circular plate and (4) four screws removed from circular plate w/ bar..

Bar

8) Insert the Ultrafine Filter onto the screen in the circular plate w/ bar. Be sure to handle the ultrafine filter with the unserrated stainless-steel forceps (Figure 2-18). Go to step 9.

Figure 2-18. Installing the ultrafilter into the ultrafine collection stage.

9) Install the circular plate onto the screen (Figure 2-20). Go to step 10

Figure 2-20. Install the Circular plate onto the ultrafine filter. Circular plate

Screen

10) Secure the circular plate with the round screws (Figure 2-21). Go to step 12.

Figure 2-21. Securing the circular plate.

11) Locate the top collection stage with the rain cover (Figure 2-22).

Figure 2-22. Inverted, top collection stage and PM- 10 PUF collection platform. Top collection stage

PM-10 PUF collection platform

Rain cover

12) Install the PM-10 PUF substrate or PM-10 grease collection platform into the top collection stage (Figures 2-23, 2-24 and 2-25).

Figure 2-23. Installing the PM-10 PUF collection platform onto one standoff. (HVCI is inverted)

Figure 2-24. Installing the PM-1 0 PUF collection platform onto the three standoffs. (HVCI is inverted)

Figure 2-25. Securing the PM-10 PUF collection platform onto the three standoffs. (HVCI is inverted)

13) Install the remaining PUF substrate collection platforms into the other collection stages. Be sure to match the correct size PUF substrate collection platform with the correct cut-point size impaction platform. For example, you should install the PM-2.5 PUF collection platform into a PM-2.5 cut-point size impaction platform. 14) Assemble the individual stages so that the largest cutpoint collection stage (PM-10 PUF or grease) is installed in the top collection stage, with the subsequent collection stages possessing smaller cut points installed underneath each other. For example, if you want to use all six possible cut point sizes (PM-10, PM-2.5, PM-1.0, PM-0.5, PM-0.1 and the ultrafilter) the PM-10 collection stage would be the top collection stage, the PM-2.5 collection stage should be installed below the PM-10 collection stage, the PM-1.0 collection stage should be installed below the PM-2.5 collection stage, the PM-0.5 collection stage should be installed below the PM-1.0 collection stage, the PM-0.1 collection stage should be installed below the PM-0.5 collection stage, and the ultrafilter collection stage should be installed below the PM-0.1 collection stage (Figure 2- 26).

Figure 2-26. Sampler assembled.

15) Determine which collection stage will be the last, or bottom, collection stage. 16) Locate the sampler base plate (Figure 2-27).

Figure 2-27. Sampler base plate.

17) Install the sampler base plate onto the bottom collection stage (Figure 2- 28). Go to Section 2.4.4 (Installing the Sampler Onto the Tripod).

Figure 2-28. Sampler base plate installed onto the bottom collection stage.

2.4.4. INSTALLING THE SAMPLER ONTO THE TRIPOD

Follow these steps to install the sampler onto the tripod:

1) Determine an appropriate location for the sampler and pump. The HVCI sampler should be located as far away from the pump’s exhaust as possible to ensure that the instrument samples a representative sample of air, and to minimize the potential for contamination. 2) Secure the tripod’s feet with spikes or screws. Spiked feet or flat feet may be used. The flat feet have 2 holes with knock-outs that can be used to secure the tripod.

3) Install the sampler onto the tripod (Figure 2-29).

Figure 2-29. Sampler installed onto the tripod.

4) Install the Model 900 Magnehelic Gage system onto the sampler base plate. (Figure 2-30).

Figure 2-30. Magnehelic Gage installed onto the base plate of the BGI900

5) Locate the air hose. 6) It is the responsibility of the end user to supply a female barbed fitting that is the correct size as the hose they will be using to connect the BGI900 to the pump. Install this fitting onto the male thread of the quick disconnect then install air hose. 7) Install the air hose onto the pump inlet. Ensure that the handles on the quick-connecting fittings are locked. Go to Section 2.4.5 (Setting Up the Pump).

2.4.5. SETTING Up THE PUMP

The pump should have a protective filter installed upstream of the blower itself. The filter will prevent soil and other foreign matter from being drawn into the system if a hose fitting detaches from the pump during sampling. The pump system (see Figure 2- 1 in section 2.1) must have an adjustable flow valve in-line that allows the operator to set the flow rate.

The flow rate is set by using a BGI High Volume Calibrator. (The latest manual can be found here: http://www.bgiusa.com/cau/manuals_2005.htm) A top inlet adaptor connects to the Model 900 and when the pump is energized the flow rate in CFM is displayed on the calibrator. The target flow rate is 31.8 CFM (900 Liters/Minute). Manually adjust the flow valve located just before the pump to set the sample flow rate at 31.8CFM. The Magnehelic Gauge is a reference of the differential pressure across the impactor. The gauge will display a number which should be recorded on the field data sheet as the calibration reference point. You should periodically check the volumetric flow through the sampler during the pump’s “break-in” period to see if this calibration reference point changes. Adjust the flow rate valve.

HiVolCal Calibrator

TOP PM-10 COLLECTION STAGE PM-2.5 PM-1.0 INTERMEDIATE COLLECTION PM-0.5 STAGES PM-0.1 EFFLUENT BOTTOM FILTER AIR COLLECTION STAGES

BYPASS TRIPOD AIR

FILTER

PUMP

Adjust the bypass valve (Figure 2-31) on the pump to match the Volumetric Flow Rate reading on the BGI Hi Vol Calibrator

Figure 2-31. Bypass valve on the pump is used to bleed air so the pump is not choked. Design sample flow rate for the HVCI is 900 L/minute, 31.8CFM.

Section 3: Post-Sampling Analysis

This section explains how to remove the PUF substrates from the sampler, and describes the substrate extraction procedures for analysis of water-soluble components and toxicological testing and the analysis procedures.

3.1. REMOVAL OF THE COLLECTION PLATFORM

After sampling is complete, you must remove the collection platforms from the collection stages. The collection platform consists of the PUF substrate installed inside the substrate holder.

Follow these steps to remove a collection platform from a collection stage:

1) Turn off the pump. 2) Disassemble the sampler by disconnecting each of the collection stages from each other. 3) Remove each collection platform from its collection stage by unscrewing the retaining nuts (Figure 3-1).

Figure 3-1. Removing the collection platform from the three standoffs.

4) Place the used collection platforms (Figure 3-2) into individual containers to prevent cross contamination. Label the containers.

Figure 3-2. PM-2.5 collection platform.

5) If you have used an ultrafilter (Figure 3-3), remove it from its retainer and place it into a suitable container that is labeled. Be sure to handle the ultrafine filter with the unserrated stainless-steel forceps (Figure 3-4).

Figure 3-3. Ultrafilter installed inside a collection stage with a spare ultrafilter.

Ultrafilter installed in collection stage

Ultrafilter

Figure 3-4. Removing the Ultrafilter from the ultrafine collection stage.

6) Wrap the containers with aluminum foils. 7) Label the outside of the aluminum foil of each container. 8) Store the containers (Section 2) until you are ready to analyze them. 9) Install an unused collection platform into each collection stage. 10) Assemble the sampler and install it onto the tripod (Section 2). 11) Turn on the pump.

3.2. SUBSTRATE EXTRACTION FOR ANALYSIS OF W-SATEROLUBLE COMPONENTS

These extraction procedures are for aqueous extraction of polyurethane foam (PUF) P1, P2, and P3 substrates and ultrafine filter (ultrafilter). These extracts are suitable for analysis of water-soluble components. Note that the volumes of the substrate sizes are not the same and the extraction solvent volume is proportional to each substrate and filter size (Figure 3-5). Because these materials cannot be wetted directly with water, methanol is used to wet the surfaces prior to aqueous extraction.

Figure 3-5. Water Substrate Water Volume volume for aqueous extraction of PUF P1 substrates and 120 ml (PM-1 0 PUF) ultrafine filter. P2 40 ml (PM-2.5)

P3 20 ml (PM-1.0, PM-0.5, PM-0.1)

Ultrafilter( 600 ml ultrafine)

T Tools and materials required for PUF substrates only: Ultrasonic bath (large enough to hold racks with culture tubes) 25 ml culture tubes with PTFE-face rubber-lined cap 50 ml culture tubes with PTFE-face rubber-lined cap 240 ml bottle beakers, medium-rounds, wide-mouth with Teflon PTFE-lined caps Disposable 1 ml plastic pipettes individually wrapped in paper/plastic Disposable 10 ml plastic pipettes individually wrapped in paper/plastic Automated pipette Tube rack(s) to hold culture tubes (must be plastic-coated or uncoated metal wire) Labels for culture tubes Glass rods Nonserrated stainless-steel forceps Stainless-steel scissors Thermometer Timer Watch glass (~ 10 cm diameter) Milli-Q grade water or equivalent Methanol

Tools and materials required for ultrafilters only: Crystallizing dish (190 mm (diameter) x 100 mm [H]) Straight-sided, round, wide-mouth flasks 1000 ml graduated cylinder Aluminum foil Parafilm

3.2.1. LABWARE W ASHING

Wash the culture tubes, stainless-steel forceps, stainless-steel scissors, crystallizing dishes, flasks, graduated cylinder, and watch glass using Liquinox detergent or equivalent; then rinse thoroughly with Milli-Q water. Cover all items with Kimwipes and allow them to dry in room air.

3.2.2. EXTRACTING PUF SUBSTRATES WITH ILLIM-Q WATER

3.2.2.1. PM-10 PUF SUBSTRATE (P1 SIZE)

Follow these steps to extract P1 PUF substrates with Milli-Q water:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Hold the P1 PUF substrate with the stainless-steel forceps. 2) Using the stainless-steel scissors, cut off 1/3 piece of the P1 PUF substrate. 3) Hold the 1/3 piece of the P1 PUF substrate with the stainless-steel forceps. 4) Using the stainless-steel scissors, cut this 1/3 piece of PUF into 32 pieces. Allow the small pieces to fall onto a watch glass. All of the pieces should be about the same size. For example, first cut the 1/3 piece in half, then in quarters, then in eighths, then in sixteenths and finally in thirty-seconds. 5) Using the stainless-steel forceps, place the 32 PUF pieces into a labeled culture tube or other suitable glass laboratory vessel. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 6) Clean the rod, forceps, scissors and watch glass with Milli-Q grade water between samples. 7) Add 6 ml of methanol to one flask using the pipette. 8) Place a cap on the flask, then gently shake to evenly wet the PUF pieces with methanol. 9) Remove the cap and, without delay (to minimize evaporation of methanol), pipette 120 ml of Milli-Q water into the culture tube (twelve 10 ml aliquots with the 10 ml pipette). 10) Use a clean, dry glass rod to disperse any small air bubbles in the PUF pieces that float near the top of the aqueous solution. 11) Replace the cap onto the top of the flask. 12) Place the flask (with the PUF pieces) in a rack. 13) Return to the original P1 PUF substrate (that has a 1/3 piece cut out) and hold it with the stainless-steel forceps. 14) Using the stainless-steel scissors, cut off another 1/3 piece of the P1 PUF substrate. 15) Repeat steps 3-12 for the second 1/3 piece of the P1 PUF substrate. 16) Hold the last remaining 1/3 piece of the original P1 PUF substrate with the stainless-steel forceps. 17) Repeat steps 2-12 for the remaining 1/3 piece of the P1 PUF substrate. 44

18) Place the rack in the ultrasonic bath. 19) Begin to sonicate the flasks starting with room temperature water. 20) Measure and record the temperature of the water in the ultrasonic bath. 21) Sonicate the flasks for 60 minutes. 22) Check the temperature of the water in the ultrasonic bath periodically. If the water temperature rises above 30º C, change the water in the bath to keep the water temperature below 30ºC. 23) Cover the flasks with Parafilm. 24) Store the covered flasks in a dark refrigerator, prior to analysis.

3.2.2.2. PM-2.5 PUF SUBSTRATE (P2 SIZE)

Follow these steps to extract P2 PUF substrates with Milli-Q water:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Hold the P2 PUF substrate with the stainless-steel forceps. 2) Using the stainless-steel scissors, cut off 1/3 piece of the P2 PUF substrate. 3) Hold the 1/3 piece of the P2 PUF substrate with the stainless-steel forceps. 4) Using the stainless-steel scissors, cut this 1/3 piece of PUF into 16 pieces. Allow the small pieces to fall onto a watch glass. All of the pieces should be about the same size. For example, first cut the 1/3 piece in half, then in quarters, then in eighths, and finally in sixteenths. 5) Using the stainless-steel forceps, place the 16 PUF pieces into a labeled, 50 ml culture tube or other suitable glass laboratory vessel. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 6) Clean the rod, forceps, scissors and watch glass with Milli-Q grade water between samples. 7) Add 2 ml of methanol to one flask using two 1 ml pipettes. 8) Place a cap on the flask, then gently shake to evenly wet the PUF pieces with methanol. 9) Remove the cap and, without delay (to minimize evaporation of methanol), pipette 40 ml of Milli-Q water into the culture tube (four 10 ml aliquots with the 10 ml pipette). 10) Use a clean, dry glass rod to disperse any small air bubbles in the PUF pieces that float near the top of the aqueous solution. 11) Replace the cap onto the top of the flask. 12) Place the flask (with the PUF pieces) in a rack. 13) Return to the original P2 PUF substrate (that has a 1/3 piece cut out) and hold it with the stainless-steel forceps. 14) Using the stainless-steel scissors, cut off another 1/3 piece of the P2 PUF substrate. 15) Repeat steps 3-12 for the second 1/3 piece of the P2 PUF substrate. 16) Hold the last remaining 1/3 piece of the original P2 PUF substrate with the stainless-steel forceps. 17) Repeat steps 2-12 for the remaining 1/3 piece of the P2 PUF substrate. 18) Place the rack in the ultrasonic bath. 19) Begin to sonicate the flasks starting with room temperature water. 20) Measure and record the temperature of the water in the ultrasonic bath. 46

21) Sonicate the flasks for 60 minutes. 22) Check the temperature of the water in the ultrasonic bath periodically. If the water temperature rises above 30º C, change the water in the bath to keep the water temperature below 30ºC. 23) Cover the flasks with Parafilm. 24) Store the covered flasks in a dark refrigerator, prior to analysis.

3.2.2.3. PM-1.0, PM-0.5 AND PM-0.1 PUF SUBSTRATE (P3 SIZE)

Follow these steps to extract P3 PUF substrates with Milli-Q water:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Hold the P3 PUF substrate with the stainless-steel forceps. 2) Using the stainless-steel scissors, cut off 1/3 piece of the P3 PUF substrate. 3) Hold the 1/3 piece of the P3 PUF substrate with the stainless-steel forceps. 4) Using the stainless-steel scissors, cut this 1/3 piece of PUF into 8 pieces. Allow the small pieces to fall onto a watch glass. All of the pieces should be about the same size. For example, first cut the 1/3 piece in half, then in quarters, and finally in eighths. 5) Using the stainless-steel forceps, place the 16 PUF pieces into a labeled, 25 ml culture tube or other suitable glass laboratory vessel. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 6) Clean the rod, forceps, scissors and watch glass with Milli-Q grade water between samples. 7) Add 1 ml of methanol to one flask using a 1 ml pipette. 8) Place a cap on the flask, then gently shake to evenly wet the PUF pieces with methanol. 9) Remove the cap and, without delay (to minimize evaporation of methanol), pipette 20 ml of Milli-Q water into the culture tube (two 10 ml aliquots with the 10 ml pipette). 10) Use a clean, dry glass rod to disperse any small air bubbles in the PUF pieces that float near the top of the aqueous solution. 11) Replace the cap onto the top of the flask. 12) Place the flask (with the PUF pieces) in a rack. 13) Return to the original P3 PUF substrate (that has a 1/3 piece cut out) and hold it with the stainless-steel forceps. 14) Using the stainless-steel scissors, cut off another 1/3 piece of the P3 PUF substrate. 15) Repeat steps 3-12 for the second 1/3 piece of the P3 PUF substrate.

16) Hold the last remaining 1/3 piece of the original P3 PUF substrate with the stainless-steel forceps. 17) Repeat steps 2-12 for the remaining 1/3 piece of the P3 PUF substrate. 18) Place the rack in the ultrasonic bath. 19) Begin to sonicate the flasks starting with room temperature water. 20) Measure and record the temperature of the water in the ultrasonic bath. 21) Sonicate the flasks for 60 minutes. 22) Check the temperature of the water in the ultrasonic bath periodically. If the water temperature rises above 30º C, change the water in the bath to keep the water temperature below 30ºC. 23) Cover the flasks with Parafilm. 24) Store the covered flasks in a dark refrigerator, prior to analysis.

3.2.2.4. ULTRAFINE FILTER (ULTRAFILTER)

Follow these steps to extract ultrafine filters with Milli-Q water:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Place the ultrafine filter inside the 190 mm crystallizing dish. 2) Hold the ultrafine filter with the stainless-steel forceps. 3) Using the stainless-steel scissors, cut the entire ultrafine filter into small pieces (about 2 cm square). 4) Place the small pieces of cut-up ultrafine filter into a straight-sided round flask. 5) Pipette 30 ml of methanol into the flask. 6) Without delay, to minimize evaporation of methanol, gently shake the flask to evenly wet the filter pieces with methanol. 7) Using the 1000 ml-graduated cylinder, immediately add 600 ml of Milli-Q water to the flask. 8) Cover the flask with a small piece of aluminum foil. 9) Place the flask directly into the water of the ultrasonic bath. 10) Begin to sonicate the flask starting with room temperature water. 11) Measure and record the temperature of the water in the ultrasonic bath. 21) Sonicate the flask for 60 minutes. 22) Check the temperature of the water in the ultrasonic bath periodically. If the water temperature rises above 30º C, change the water in the bath to keep the water temperature below 30ºC. 23) Cover the flask with Parafilm. 24) Store the covered flask in a dark refrigerator, prior to analysis.

3.3. SUBSTRATE EXTRACTION FOR TOXICOLOGICAL TESTING

These extraction procedures are for solvent extraction of polyurethane foam (PUF) P1, P2, and P3 substrates and ultrafine filters (ultrafilter). These extracts are suitable for analysis of water-soluble components. Note that the volumes of the substrate sizes are not the same and the extraction solvent volume is proportional to each substrate and filter size (Figure 3-4).

Tools and materials required for PUF substrates only: Laboratory exhaust hood Ultrasonic bath (large enough to hold a rack with the extraction tubes) 25 ml culture (“extraction”) tubes with PTFE-face rubber-lined caps 40 ml (“evaporation”) tubes with Teflon-lined, solid top cap 4 ml (“storage”) glass vials with PTFE Teflon-lined caps Pasteur pipettes (9 inch) Evaporator (ORGANOMATION N-VAP model 111 or equivalent) Ultra-High Purity grade nitrogen gas cylinder (99.99%) Regulator for nitrogen gas Tube rack to hold extraction tubes Balance (Mettler AE 163, sensitivity ± 0.01 mg or equivalent) Labels for tubes and vials Disposable 1 ml plastic pipettes individually wrapped in paper/plastic Disposable 10 ml plastic pipettes individually wrapped in paper/plastic Automated pipette Tube rack(s) to hold culture tubes (plastic-coated or uncoated metal wire) Glass rods Nonserrated stainless-steel forceps Stainless-steel scissors Thermometer Timer Watch glass (~ 10 cm diameter) Milli-Q grade water or equivalent Methanol Aluminum foil

Tools and materials required for ultrafilters only: Crystallizing dish (190 mm (diameter) x 100 mm [H]) 250 ml Erlenmeyer (“extraction”) flasks 500 ml Erlenmeyer (“storage”) flasks 250 ml graduated cylinder

3.3.1. LABWARE W ASHING

3.3.2. EXTRACTING PUF SUBSTRATES WITH METHANOL

3.3.2.1. PM-10 PUF SUBSTRATE (P1 SIZE)

Follow these steps to extract P1 PUF substrates with methanol:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Hold the P1 PUF substrate with the stainless-steel forceps. 2) Using the stainless-steel scissors, cut off 1/3 piece of the P1 PUF substrate. 3) Hold the 1/3 piece of the P1 PUF substrate with the stainless-steel forceps. 4) Using the stainless-steel scissors, cut this 1/3 piece of PUF into 32 pieces. Allow the small pieces to fall onto a watch glass. All of the pieces should be about the same size. For example, first cut the 1/3 piece in half, then in quarters, then in eighths, then in sixteenths and finally in thirty-seconds. 5) Using the stainless-steel forceps, place the 32 PUF pieces into a 250 ml extraction flask. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 6) Clean the rod, forceps, scissors and watch glass with Milli-Q grade water between samples. 7) Add 15 ml of methanol to the flask using the automatic pipette and a 10 ml pipette. 8) Place a cap on the flask, then gently shake to evenly wet the PUF pieces with methanol. 9) Place the flask (with the PUF pieces) in a rack. 10) Return to the original P1 PUF substrate (that has a 1/3 piece cut out) and hold it with the stainless-steel forceps. 11) Using the stainless-steel scissors, cut off another 1/3 piece of the P3 PUF substrate. 12) Repeat steps 3-9 for the second 1/3 piece of the P3 PUF substrate. 13) Hold the last remaining 1/3 piece of the original P3 PUF substrate with the stainless-steel forceps. 14) Repeat steps 2-9 for the remaining 1/3 piece of the P3 PUF substrate. 15) Place the rack in the ultrasonic bath. 16) Begin to sonicate the flasks starting with room temperature water. 17) Measure and record the temperature of the water in the ultrasonic bath. 18) Sonicate the flasks for 60 minutes. 19) Check the temperature of the water in the ultrasonic bath periodically. If the water temperature rises above 30º C, change the water in the 53

20) Use a 10 ml pipette to transfer the methanol extracts from the extraction tubes to labeled 40 ml evaporation tubes. 21) Cap the extraction tubes and evaporation tubes except during transfers. 22) Transfer as much extract from each tube as is conveniently possible. 23) Record the approximate amount of solvent transferred using the 10 ml pipette. 24) Add 10 ml of fresh methanol to each extraction tube. 25) Place the extraction tubes in a rack. 26) Repeat steps 15-19. 27) Repeat steps 20-23. 28) Add 10 ml of fresh methanol to each extraction tube. 29) Place the extraction tubes in a rack. 30) Repeat steps 15-19. 31) Repeat steps 20-23. The approximate overall efficiency of extraction can be calculated using measured amounts of methanol added and approximate amounts of extract transferred for each of the three extractions. For example, you started with 15 ml for original extraction, transferred most to evaporation tube, added 10 ml to the extraction tubes, sonicated, and transferred most to the evaporation tubes, added second 10 ml to extraction tubes, son icated, and transferred most of the extract to the evaporation tube.

3.3.2.2. PM-2.5 PUF SUBSTRATE (P2 SIZE)

Follow these steps to extract P2 PUF substrates with methanol:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Hold the P2 PUF substrate with the stainless-steel forceps. 2) Using the stainless-steel scissors, cut off 1/3 piece of the P2 PUF substrate. 3) Hold the 1/3 piece of the P2 PUF substrate with the stainless-steel forceps. 4) Using the stainless-steel scissors, cut this 1/3 piece of PUF into 16 pieces. Allow the small pieces to fall onto a watch glass. All of the pieces should be about the same size. For example, first cut the 1/3 piece in half, then in quarters, then in eighths, and finally in sixteenths. 5) Using the stainless-steel forceps, place the 16 PUF pieces into a labeled, 50 ml culture tube or other suitable glass laboratory vessel. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 5) Using the stainless-steel forceps, place the 16 PUF pieces into a 250 ml extraction flask. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 6) Clean the rod, forceps, scissors and watch glass with Milli-Q grade water between samples. 7) Add 15 ml of methanol to the flask using the automatic pipette and a 10 ml pipette. 8) Place a cap on the flask, then gently shake to evenly wet the PUF pieces with methanol. 9) Place the flask (with the PUF pieces) in a rack. 10) Return to the original P2 PUF substrate (that has a 1/3 piece cut out) and hold it with the stainless-steel forceps. 11) Using the stainless-steel scissors, cut off another 1/3 piece of the P3 PUF substrate. 12) Repeat steps 3-9 for the second 1/3 piece of the P3 PUF substrate. 13) Hold the last remaining 1/3 piece of the original P3 PUF substrate with the stainless-steel forceps. 14) Repeat steps 2-9 for the remaining 1/3 piece of the P3 PUF substrate. 15) Place the rack in the ultrasonic bath. 16) Begin to sonicate the flasks starting with room temperature water. 17) Measure and record the temperature of the water in the ultrasonic bath. 18) Sonicate the flasks for 60 minutes. 19) Check the temperature of the water in the ultrasonic bath periodically. If 55

the water temperature rises above 30º C, change the water in the bath to keep the water temperature below 30ºC. 20) Use a 10 ml pipette to transfer the methanol extracts from the extraction tubes to labeled 40 ml evaporation tubes. 21) Cap the extraction tubes and evaporation tubes except during transfers. 22) Transfer as much extract from each tube as is conveniently possible. 23) Record the approximate amount of solvent transferred using the 10 ml pipette. 24) Add 10 ml of fresh methanol to each extraction tube. 25) Place the extraction tubes in a rack. 26) Repeat steps 15-19. 27) Repeat steps 20-23. 28) Add 10 ml of fresh methanol to each extraction tube. 29) Place the extraction tubes in a rack. 30) Repeat steps 15-19. 31) Repeat steps 20-23. The approximate overall efficiency of extraction can be calculated using measured amounts of methanol added and approximate amounts of extract transferred for each of the three extractions. For example, you started with 15 ml for original extraction, transferred most to evaporation tube, added 10 ml to the extraction tubes, sonicated, and transferred most to the evaporation tubes, added second 10 ml to extraction tubes, son icated, and transferred most of the extract to the evaporation tube.

3.3.2.3. PM-1.0, PM-0.5 AND PM-0.1 PUF SUBSTRATE (P3 SIZE)

Follow these steps to extract P3 PUF substrates with methanol:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Hold the P3 PUF substrate with the stainless-steel forceps. 2) Using the stainless-steel scissors, cut off 1/3 piece of the P3 PUF substrate. 3) Hold the 1/3 piece of the P3 PUF substrate with the stainless-steel forceps. 4) Using the stainless-steel scissors, cut this 1/3 piece of PUF into 8 pieces. Allow the small pieces to fall onto a watch glass. All of the pieces should be about the same size. For example, first cut the 1/3 piece in half, then in quarters, and finally in eighths. 5) Using the stainless-steel forceps, place the 8 PUF pieces into a labeled, 50 ml culture tube or other suitable glass laboratory vessel. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 5) Using the stainless-steel forceps, place the 8 PUF pieces into a 250 ml extraction flask. If necessary, use a glass rod to push the PUF pieces down to the bottom of the extraction flask. 6) Clean the rod, forceps, scissors and watch glass with Milli-Q grade water between samples. 7) Add 15 ml of methanol to the flask using the automatic pipette and a 10 ml pipette. 8) Place a cap on the flask, then gently shake to evenly wet the PUF pieces with methanol. 9) Place the flask (with the PUF pieces) in a rack. 10) Return to the original P3 PUF substrate (that has a 1/3 piece cut out) and hold it with the stainless-steel forceps. 11) Using the stainless-steel scissors, cut off another 1/3 piece of the P3 PUF substrate. 12) Repeat steps 3-9 for the second 1/3 piece of the P3 PUF substrate. 13) Hold the last remaining 1/3 piece of the original P3 PUF substrate with the stainless-steel forceps. 14) Repeat steps 2-9 for the remaining 1/3 piece of the P3 PUF substrate. 15) Place the rack in the ultrasonic bath. 16) Begin to sonicate the flasks starting with room temperature water. 17) Measure and record the temperature of the water in the ultrasonic bath. 18) Sonicate the flasks for 60 minutes. 19) Check the temperature of the water in the ultrasonic bath periodically. If 57

3.3.3. EVAPORATION OF SOLVENT FROM PUF EXTRACTS

Follow these steps to evaporate solvent from PUF substrates:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Remove the caps from as many evaporation tubes as will fit into the evaporator. 2) Place the evaporation tubes into the evaporator. If the evaporator has a provision for the use of a water bath, maintain the bath temperature at 30° C. 3) Place clean Pasteur pipettes in the evaporator. 4) Cover the opening of each evaporation tube with a piece of aluminum foil. 5) Open the valves of the pipettes and bring the tip of each pipette close to the solvent surface (inside the evaporation tubes) until there is a very slight indenture in the surface due to the gas flow. 6) As the methanol evaporates, add fresh methanol to wash the walls of the evaporation tubes. 7) Evaporate until the volume in the evaporation tubes is less than 3 ml. 8) Remove the evaporation tubes from the evaporator. 9) Cap each evaporation tube until you are ready to transfer the concentrated extract to the storage tubes. Transfer of extract to the storage tubes 10) Use a clean Pasteur pipette to transfer the concentrated extract from the evaporation tubes to preweighed, labeled storage tubes. Be sure to weigh the storage tubes with their caps on. The cap used for weighing must be identifiable to make sure that the same cap is used for post-weighing of each tube. 11) Replace the caps on the evaporation tubes until you are ready to rinse them into the storage tubes. 12) Replace the caps on the storage tubes until you are ready to evaporate the concentrated extract. 13) Remove the caps from as many storage tubes as will fit into the evaporator.

14) Place the storage tubes in the evaporator. 15) Evaporate most of the concentrated extract in the storage tubes. First rinse of the evaporation tubes 16) Using a 1 ml pipette, add 0.5 ml of fresh methanol to rinse the evaporation tubes. 17) Transfer this first 0.5 ml rinse of the evaporation tubes to the storage tubes. 18) Repeat steps 4-8. 19) Evaporate most of the concentrated extract in the storage tubes. Second rinse of the evaporation tubes 20) Using a 1 ml pipette, add another 0.5 ml of fresh methanol to rinse the evaporation tubes a second time. 21) Transfer this second 0.5 ml rinse of the evaporation tubes to the storage tubes. 22) Repeat steps 4-8. 23) Evaporate most of the concentrated extract in the storage tube. Third rinse of the evaporation tubes 24) Using a 1 ml pipette, add another 0.5 ml of fresh methanol to rinse the evaporation tubes a third time. 25) Transfer this third 0.5 ml rinse of the evaporation tubes to the storage tubes. 26) Repeat steps 4-8. 27) Evaporate the concentrated extract until it is dry. 28) Replace the caps on the storage tubes. Ensure that each cap corresponds to the original pre-weighing of each storage tube. 29) Weigh each storage tube and record its weight. 30) Calculate the weight of the extract by subtracting the original weight of the storage tube from the final weight of the storage tube. Repeat this for each storage tube and record all values. 31) Place the storage tubes in a freezer at -20° C until you are ready to begin the toxicological analysis (Section 3.4).

3.3.4. EXTRACTING ULTRAFILTER FILTERS WITH METHANOL

Follow these steps to extract ultrafilter filters with methanol:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Place the ultrafine filter inside the 190 mm crystallizing dish. 2) Hold the ultrafine filter with the stainless-steel forceps. 3) Using the stainless-steel scissors, cut the entire ultrafine filter into small pieces (about 2 cm square). 4) Place the small pieces of cut-up ultrafine filter into a 250 ml Erlenmeyer (“extraction”) flask. 5) Using the graduated cylinder, add 120 ml of methanol into the extraction flask. 6) Use a small piece of aluminum foil to cover the flask. 7) Place the flask directly into the water of the ultrasonic bath. 8) Begin to sonicate the flask starting with room temperature water. 9) Measure and record the temperature of the water in the ultrasonic bath. 10) Sonicate the flask for 60 minutes. 11) Check the temperature of the water in the ultrasonic bath periodically. If the water temperature rises above 30º C, change the water in the bath to keep the water temperature below 30ºC. 12) Pour this first methanol extract from the original extraction flask into another clean 500 ml Erlenmeyer (“storage”) flask. 13) Add an additional 120 ml of fresh methanol to the original extraction flask. 14) Use a small piece of aluminum foil to cover the original extraction flask. 15) Place the original extraction flask directly into the water of the ultrasonic bath. 16) Repeat steps 8-11. 17) Pour the second methanol extract into the same storage flask used in step 12.

18) Add an additional 120 ml of fresh methanol to the original extraction flask. 19) Use a small piece of aluminum foil to cover the original extraction flask. 20) Place the original extraction flask directly into the water of the ultrasonic bath. 21) Repeat steps 8-11. 22) Pour the third methanol extract into the same storage flask used in step 12. 23) Cap the 500 ml Erlenmeyer storage flasks containing the extracts until you are ready to begin the evaporation procedure (Section 3.3.5).

Note that the total amount of methanol retained by the ultrafine filter is about 30 ml. Thus the first aliquot of methanol transferred from the extraction flask contains about 75% (90/120) of the material dissolved in the solvent. The second aliquot of methanol transferred from the extraction flask contains about 80% (120/150) of the remaining material (25%). The third aliquot transferred contains about 80% (120/150) of the remaining material (5%). Therefore, the total fraction transferred from the extraction flask is about (75 + 0.8(25) + 0.8(5))% = 99%.

3.3.5. EVAPORATION OF SOLVENT FROM ULTRAFINE EXTRACTS

Follow these steps to evaporate solvent from ultrafine extracts:

NOTE: It is important to wear powder-free gloves at all times. Rinse the gloves with Milli-Q water and dry the gloves by wiping them with Kimwipes. 1) Determine the maximum number (N) of evaporation tubes that can be used simultaneously with the evaporator. 2) Remove the caps of the evaporation tubes. Transfer of extract to each of the “N” evaporation tubes 3) Using a pipette, transfer 35 ml of extract from the storage flask to each of the “N” evaporation tubes. 4) Place the evaporation tubes into the evaporator. If the evaporator has a provision for the use of a water bath, maintain the bath temperature at 30° C. 5) Place clean Pasteur pipettes in the evaporator. 6) Cover the opening of each evaporation tube with a piece of aluminum foil. 7) Open the valves of the pipettes and bring the tip of each pipette close to the solvent surface (inside the evaporation tubes) until there is a very slight indenture in the surface due to the gas flow. 8) As the methanol evaporates, add fresh methanol to wash the walls of the evaporation tubes. 9) Continue transferring until all the extract has been removed from the storage flask (step 3). First rinse of the storage flask 10) After all of the extract has been removed from the storage flask, add 25 ml of fresh methanol to rinse the storage flask. 11) Transfer 35 ml of the first rinse (using a pipette) from the storage flask to each of the “N” evaporation tubes. 12) Repeat steps 4-8. 13) Continue transferring until all the first rinse has been removed from the storage flask (step 3).

Second rinse of the storage flask 14) After all of the first rinse has been removed from the storage flask, add 25 ml of fresh methanol to rinse the storage flask a second time. 15) Transfer 35 ml of the second rinse (using a pipette) from the storage flask to each of the “N” evaporation tubes. 16) Repeat steps 4-8. 17) Continue transferring until all the second rinse has been removed from the storage flask (step 3). 18) Evaporate until volume is less than 3 ml. Transfer of extract to “Tube A” and “Tube B” 19) Using a pipette, transfer all of the remaining extract from the “N” evaporation tubes to only two of the evaporation tubes (“Tube A” and “Tube B”). 20) Repeat steps 4-8. 21) Evaporate the extract in Tube A and Tube B until the volume is less than 3 ml in each tube. First rinse of the “N” evaporation tubes 22) Using a 1 ml pipette, add 1 ml of fresh methanol to rinse the other “N” evaporation tubes. 23) Transfer this first 1 ml rinse of the other “N” evaporation tubes to Tube A and Tube B (step 19). 24) Repeat steps 4-8. 25) Evaporate the first 1 ml rinse until the volume is less than 3 ml in each tube. Second rinse of the “N” evaporation tubes 26) Using a 1 ml pipette, add another 1 ml of fresh methanol to rinse the other “N”evaporation tubes a second time. 27) Transfer this second 1 ml rinse of the other “N” evaporation tubes to Tube A and Tube B (step 19). 28) Repeat steps 4-8. 29) Evaporate the second 1 ml rinse until the volume is less than 3 ml in each tube.

Third rinse of the “N” evaporation tubes 30) Using a 1 ml pipette, add another 1 ml of fresh methanol to rinse the other “N” evaporation tubes a third time. 31) Transfer this third 1 ml rinse of the other “N” evaporation tubes to Tube A and Tube B (step 19). 32) Repeat steps 4-8. 33) Evaporate the third 1 ml rinse until the volume is less than 3 ml in each tube. Transfer of extract to “Tube B” 34) Using a pipette, transfer the extract from one of the two remaining evaporation tubes (Tube A) to the other evaporation tube (Tube B). 35) Repeat steps 4-8. 36) Evaporate the extract in Tube B until the volume is less than 3 ml. First rinse of “Tube A” 37) Using a 1 ml pipette, add 1 ml of fresh methanol to rinse Tube A. 38) Transfer this first 1 ml rinse of Tube A to Tube B. 39) Repeat steps 4-8. 40) Evaporate the first 1 ml rinse until the volume is less than 3 ml in Tube B. Second rinse of “Tube A” 41) Using a 1 ml pipette, add another 1 ml of fresh methanol to rinse Tube A a second time. 42) Transfer this second 1 ml rinse of Tube A to Tube B. 43) Repeat steps 4-8. 44) Evaporate the second 1 ml rinse until the volume is less than 3 ml in Tube B. Third rinse of “Tube A” 45) Using a 1 ml pipette, add another 1 ml of fresh methanol to rinse Tube A a third time. 46) Transfer this third 1 ml rinse of Tube A to Tube B. 47) Repeat steps 4-8. 48) Evaporate the third 1 ml rinse until the volume is less than 3 ml in Tube B. 49) Remove Tube B from the evaporator. 50) Cap Tube B until you are ready to transfer the concentrated extract to the storage tube. Transfer of extract to the storage tube 51) Use a clean Pasteur pipette to transfer the concentrated extract from Tube B to a preweighed, labeled storage tube. Be sure to weigh the storage tube with its cap on. The cap used for weighing must be 65

identifiable to make sure that the same cap is used for post-weighing of the storage tube. 52) Replace the cap on Tube B until you are ready to rinse it into the storage tube. 53) Replace the cap on the storage tube until you are ready to evaporate the concentrated extract. 54) Remove the cap from the storage tube. 55) Place the storage tube in the evaporator. 56) Evaporate most of the concentrated extract in the storage tube. First rinse of the storage tube 57) Using a 1 ml pipette, add 0.5 ml of fresh methanol to rinse Tube B. 58) Transfer this first 0.5 ml rinse of Tube B to the storage tube. 59) Repeat steps 4-8. 60) Evaporate most of the concentrated extract in the storage tube. Second rinse of the storage tube 61) Using a 1 ml pipette, add another 0.5 ml of fresh methanol to rinse Tube B a second time. 62) Transfer this second 0.5 ml rinse of Tube B to the storage tube. 63) Repeat steps 4-8. 64) Evaporate most of the concentrated extract in the storage tube. Third rinse of the storage tube 65) Using a 1 ml pipette, add another 0.5 ml of fresh methanol to rinse Tube B a third time. 66) Transfer this third 0.5 ml rinse of Tube B to the storage tube. 67) Replace the cap on the storage tube. Ensure that the cap corresponds to the original pre-weighing of the storage tube. 68) Weigh the storage tube and record its weight. 72) Calculate the weight of the extract by subtracting the original weight of the storage tube from the final weight of the storage tube. 73) Place the storage tube in a freezer at -20° C until you are ready to begin the toxicological analysis (Section 3.4).

3.4. ANALYSIS OF SAMPLES

The Model 900 HVCI samples can be analyzed for Toxicological, Biological, Chemical Speciation and Radionuclide Analysis:

• Water-soluble components such as sulfate, nitrate and major cations, with ion chromatography • Light organic acids such as formic and acetic acids, with ion chromatography • Low molecular weight acids, such as adipic acid, hexanoic acid, with gas chromatography coupled with a flame-ionization detector • Aldehydes and other functional organic compounds, with high-performance liquid chromatography • Polynuclear aromatic hydrocarbons, with gas chromatography coupled with mass spectrometry, or high-performance liquid chromatography coupled with a fluorescent detector • Polychlorinated biphenyls and dioxins, using gas chromatography coupled with tandem mass spectrometry • Trace metals, using induced conductivity plasma coupled with mass spectrometry • Total organic and elemental carbon, using thermal optical reflectance • Strong acidity, with a pH-meter • Macromolecules, with pyrolysis-gas chromatography-mass spectrometry • Aerosolization of aqueous suspension, or pre-collected particles, using nebulizers • The size and shape of particles, using light microscopy, or electron imaging • Molecular characterization, using Fourier-Transform Infrared Spectros- copy • A wide variety of biological and toxicological tests. • Radionuclide using laboratory Alpha, Beta and Gamma Detectors.

Reference Publications:

P. Demokritou, I.G. Kavouras, S. T. Ferguson, & P. Koutrakis: “Development of a High Volume Cascade Impactor for Toxicological and Chemical Characterization Studies”, Aerosol Science & Technology 36: 925-933 (2002)

United States Patent 6,435,043 Ferguson , et al. August 20, 2002

Impaction substrate and methods of use. The present disclosure relates to a method of collecting particles in a gas sample comprising impacting particles in the gas sample on a porous material, as well as to impactors wherein an inertial impactor includes a porous substrate.

Inventors: Ferguson; Stephen T. (N. Billerica, MA), Kavouras; Ilias G. (Boston, MA), Wolfson; Jack M. (Jamaica Plain, MA), Koutrakis; Petros (Wellesley, MA)