Inhalation exposure to three-dimensional printer emissions stimulates acute hypertension and microvascular dysfunction.

Animal Model

All procedures and experiments in this study conformed to the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals (8th Edition) and were approved by the West Virginia University Animal Care and Use Committee. Male, Sprague-Dawley rats (7–8 weeks of age) were purchased from Hilltop Laboratories (Scottdale, PA), and housed in an AAALAC approved animal facility at the West Virginia University Health Sciences Center. All animals were maintained on a 12-hour light/dark cycle, provided food and water ad libitum , and acclimated for at least 72 hours prior to training and/or use. Animals were then randomly assigned to either the Sham-Control (filtered air) or 3DPE inhalation exposure groups. 3DPE Generation and Aerosol Inhalation Exposure All rats were progressively trained daily for ~2 weeks prior to nose-only exposures. During these sessions, rats were placed in restraining tubes (Allay ® , DSI, St. Paul, MN) initially for ~5 minutes, and progressively increased in 15–20 minutes intervals for up to 4 hours. Tubes were wrapped in red polyurethane sheets to reduce visual stress. At any point during training or exposure, if rats displayed distress, they were immediately removed. On experimental days, rats were placed in a nose-only exposure system (Inhalation Tower, DSI, St. Paul, MN) for 3–4 hours. The stainless steel, nose-only exposure device is designed to accommodate up to 14 rodents. The tower has two levels of seven exposure ports, radially positioned around the aerosol delivery components of the system. A desktop 3D printer (Replicator ® 2x, MakerBot Industries, Brooklyn, NY) was placed in a 500 L stainless steel chamber. Black ABS is among the more common filaments used in 3D printing, and was used for all rat inhalation exposures herein. The printer was operated continuously during rat exposures. Emissions from the chamber were pumped into the nose-only exposure tower. Real-time and time-integrated aerosol sampling and monitoring instrumentation analyzed the emissions in the generation chamber and nose-only exposure tower as previously described ( Yi et al. , 2013 ). Briefly, 3DPE particle size distribution and mass concentration were measured continuously with a scanning mobility particle sizer (SMPS, TSI Inc, Shoreview, MN), and an electric low-pressure impactor (ELPI, Dekati, Ltd., Kangasala, Finland). The aerosol mass concentration was verified gravimetrically. Intravital Microscopy At 24 hr. post-exposure, rats were anesthetized with thiobutabarbital sodium (Inactin, 100 mg/kg, i.p.), placed on a heating plate, connected to a thermocouple probe to maintain a 37°C rectal temperature with an Animal Temperature Controller (World Precision Instruments, Sarasota, Florida). The trachea was intubated to ensure a patent airway, and the right carotid artery was cannulated to measure arterial pressure. The right spinotrapezius muscle was then exteriorized for microscopic observation, leaving its innervation and all feed vessels intact. After exteriorization, the muscle was gently secured over an optical pedestal at its in situ length. The muscle was next enclosed in a tissue bath for transillumination and observation. Throughout the surgery and all experimental periods, the muscle was continuously superfused with an electrolyte solution (119 mM NaCl, 25 mM NaHCO 3 , 6 mM KCl and 3.6 mM CaCl 2 ), warmed to 35°C, and equilibrated with 95% N 2 - 5% CO 2 (pH=7.35–7.40). Superfusate flow rate was maintained at 4–6 mL/min to minimize equilibration with atmospheric oxygen ( Boegehold and Bohlen, 1988 ). The animal preparation was then transferred to the stage of an intravital microscope, coupled to a CCD color video camera (BX51WI and DP71, respectively, Olympus, Tokyo, Japan). Observations were made with a 20X water immersion objective (final video image magnification = 1460X). One to three arterioles were studied per rat. Real-time images were displayed on a high-definition computer monitor and digitally captured for off-line analysis ( Nurkiewicz et al. , 2008 ; Nurkiewicz et al. , 2009 ). Arteriolar inner diameters were measured with Image-J software (National Institutes of Health, Bethesda, MD) calibrated with a stage micrometer. Microiontophoresis Micropipettes were custom fabricated with a Flaming/Brown Micropipette Puller (P-97, Sutter Instruments, Novato, CA). Aluminosilicate glass capillary tubes (A100-64-10, Sutter Instruments) were pulled to an inner diameter of 2–4 μm and subsequently double-beveled (BV-10, Sutter Instruments), as previously described ( Nurkiewicz and Boegehold, 2004 ). Micropipettes were then backfilled with the muscarinic agonist acetylcholine (0.025 M; ACh), or the nitric oxide donor sodium nitroprusside (0.05 M; SNP). Backloaded pipettes were attached to a microelectrode holder with an indwelling Ag/AgCl wire that was connected to a Dual Microiontophoresis Current Programmer (SYS-260; World Precision Instruments, Sarasota, FL). The microelectrode holder was attached to a three-axis hydraulic micromanipulator, combined with a one-axis hydraulic micromanipulator (MMO-203, and MMO-220A respectively, Narishige, Tokyo, Japan) to enable four-dimensional controlled movements. The entire apparatus was mounted on a motorized stage platform (GMHB-BX, Gibraltar Industries, Buffalo, NY), adjacent to the animal preparation. Finally, an additional Ag wire was submerged in the intravital tissue bath to complete the electrical circuit. The micropipette was positioned with the tip within the arteriolar adventitial layer, slightly superior to the vessel wall to prevent accidental puncture associated with tissue movement and dilation. Holding currents of 200–500 nA were used to contain agonists in the micropipette during all control and recovery periods. Microiontophoretic ejection currents were randomly generated to evaluate endothelium-dependent dilation (ACh; 20, 40, and 100 nA), and endothelium-independent dilation (SNP; 5, 10 and 20 nA). At the end of all experiments, passive maximum arteriolar diameter was established by superfusing the tissue with 10 −4 M adenosine (ADO). Formulas, Data and Statistical Analysis Mean arterial pressure (MAP) was calculated as: MAP = diastolic pressure + (systolic pressure – diastolic pressure)/3 . Arteriolar diameter (D, μm) was sampled at 10-second intervals during all control and ejection periods. Resting vascular tone was calculated for each vessel as follows: Tone = [(D pass − D c )/D pass ] × 100 , where D pass is passive diameter under ADO and D c is the diameter measured during the control period (resting diameter). A tone of 100% represents complete vessel closure, and 0% represents the passive state. To evaluate arteriolar responsiveness between individual groups with subtle differences in resting diameter, arteriolar diameter was normalized. In this case, arteriolar diameter was expressed as a percent change from control and was calculated for each vessel as follows: Diameter (% change from control) = [(D SS − D c ) - 1 × 100] , where D SS is the steady state diameter achieved during ejection. All data are reported as means ± SE, where “N” represents the number of rats studied and “n” represents the number of arterioles evaluated. One to three microvessels were studied per rat. Statistical analysis was performed by commercially available software (Sigmaplot, Systat Software Inc., San Jose, CA). One-way repeated measures ANOVA was used to determine the effect of a treatment within a group, or differences among groups. Two-way repeated measures ANOVA was used to determine the effects of group, treatment and group-treatment interactions on measured variables. For all ANOVA procedures, the Student-Newman-Keuls method for post-hoc analysis was used to isolate pairwise differences among specific groups. Significance was assessed at the 95% confidence level (P < 0.05) for all tests.