Use of conventional cigarette smoking and e-cigarette vaping for experimental stroke studies in mice

Abstract

Cigarette smoking is a major prodromal factor for the onset of many adverse health effects that may occur in the short run and is the leading cause of preventable disease, disability, and death in the United States. Moreover, it is well established that chronic smoking is associated with vascular endothelial dysfunction in a causative and dose-dependent manner primarily related to the release of reactive oxygen species (ROS), nicotine, and the induction of oxidative stress (OS) -driven inflammation [1]. Preclinical studies have also shown that nicotine (the principal e-liquid ingredient used in e-cigarettes) can also cause OS, exacerbating cerebral ischemia and secondary brain injury. Likewise, chronic e-Cig vaping could be prodromal to cerebrovascular impairment and promote cerebrovascular conditions favoring stroke onset and worsening post-ischemic brain injury. Therefore, using mice models is crucial to understand how xenobiotics such as those released by conventional and/or E-cigs can impact the onset and severity of stroke. To appropriately model human-like smoking/vaping behavior in mice, however, the exposure to these xenobiotics must be standardized and undertaken in a controlled environment. This chapter describes a well-validated protocol to reproduce standardized chronic tobacco smoke or e-cigarette vape exposure in mice in the setting of a transient ischemic stroke procedure.

Introduction

Chronic smoking is accountable for over 480,000 deaths each year in the United States (US). It is well established that active and passive cigarette smoking (CS) has been shown to nearly double the risk for stroke, stroke recurrence, and worsened outcomes [2–5] by promoting vascular endothelial dysfunction in a causative and dose-dependent manner [6]. One of the primary reasons is related to the smoke content of reactive oxygen species (ROS), nicotine, and oxidative stress (OS) -driven inflammation [1, 7–9]. Likewise, chronic e-cigarettes (e-Cig) vaping, which has become the sought-after product [10, 11] and is rapidly gaining consumers among adults and adolescents, also promotes cerebrovascular impairment setting a favorable condition for the onset of stroke and/or worsening post-ischemic outcome that is not dissimilar from that of conventional cigarettes [12]. Overall, the stroke risk factor for smokers during their lifetime is more than double that of never-smokers, and meta-analysis suggests that the risk is even greater for women based on ethnicity [13]. Even upon smoking cessation, former chronic smokers remain at considerable risk for stroke for several years [14].

Exposure to cigarette smoke has been shown to augment pro-inflammatory cytokines such as TNF-alpha, IL-1, IL-8, and GM-CSF [15] and affect multiple organ systems, thus resulting in numerous tobacco-induced diseases [15]. Burning Cigarettes can produce as many as 7000 different components. While the carcinogenic effects of cigarette smoke have been well characterized, the inflammatory and immunomodulatory effects of these various chemicals, particularly those generated by e-cig vaping, are much less defined [16]. Therefore, it is important to develop viable in vivo testing in translationally relevant models.

In the procedures described below, test animals are exposed to either CS or e-Cig vapors 6–8 times a day (2 cigarettes/hour, 6 hours/day, 7 days/week for 4 weeks) to reproduce a status of chronic exposure simulating a generalized human smoking behavior. Control animals are instead exposed to HEPA-filtered air to minimize the likelihood of any contaminants.

To generate sidestream smoke/vape, we use a computer-controlled Cigarette Smoking Machine (from CH Technologies, Westwood, NJ) which can be programmed to generate consistent, reproducible puff volumes and durations consistent with the Federal Trade Commission (FTC) smoking protocol developed based on cursory observations of human smoking behavior [16]. The standard conditions and parameters for the routine analytical generation and collection of aerosol from e-cigarettes are instead specified in the CORESTA protocol [17], which we also described in this chapter.

In summary, test animals are chronically exposed to either CS or e-Cig vapors 6–8 times a day (2 cigarettes/hour, 6 hours/day, 7 days/week for 4 weeks).

Exposure to CS is achieved by burning conventional standardized cigarettes ranging from full flavor (e.g.,3R4F reference cigarettes) to light cigarettes (1R5F reference cigarettes) available through the University of Kentucky, Tobacco Research Institute. Exposure to e-cig vapor instead is achieved using Standardized Research Electronic Cigarette (SREC) for Clinical Research or commercial products (e.g., vaping BLU ^™). Using dedicated Cigarette Smoking machines is recommended to eliminate the risk of cross-contamination between CS and e-Cigs. Note that the Cigarette Smoking machines will produce 100% sidestream smoke/vape.

Ischemic stroke is induced via transient middle cerebral artery occlusion (tMCAO) using a modified suture filament to temporarily (30 to 120 minutes) occlude blood flow through the cerebral artery.

2. Material (for chronic smoking/vaping)

2.1. Animals

6- to 8-month-old and a bodyweight comprised between 20–22 g, male and female C57BL/6 mice (see Note 1 and 2).

Before initiating the cycles of exposure to conventional or electronic cigarettes, animal body weight needs to be measured (and then monitored every 4 to 5 days). Also, a blood sample (from the tail vein) must be collected to assess the baseline parameters, including glucose levels and any marker of interest. Like body weight, blood samples need to be collected regularly when the body weight is checked out to monitor the glucose levels and markers of interest. Measurements of nicotine and cotinine levels are also useful to assess the level and consistency of the exposure between animals (see Note 3)

2.2. Conventional and E-cigarettes

Regarding conventional cigarettes, in addition to commercial products, the user can choose between different available nicotine research cigarette types (see Note 4). Concerning the E-cigarettes, the standardized products are available directly from NJOY LLC (see Note 5)

2.3. Equipment

• CSM-SCSM Cigarette Smoking Machine (CH Technologies, Westwood, NJ)

• Chemical hood

• Gas induction chamber

• Oxygen tank

• Pressure regulator

• Tubing & connectors

3. Methods

Generation of cigarette smoke and chronic animal exposure

1. The smoking machine and the induction chambers must reside inside a chemical hood with proper ventilation (see Fig. 1A).

2. To set up the system, connect the gas induction chamber to CSM-SCSM Cigarette Smoking Machine (CH Technologies, Westwood, NJ) outlet and the oxygen supply using a T connector. Oxygenated air needs to be supplied (Oxygen rate of 0.5–1%). This prevents mice from asphyxiating due to the exclusive ventilation originating from the smoking machine outlet.

3. Place the mice in the gas induction chamber (see Fig. 1B). Up to 4 mice can be exposed to smoke simultaneously using two airtight chambers.

4. Attach the cigarette (conventional or e-cigarette) to the CSM-SCSM Cigarette Smoking Machine inlet using the proper adapter. Place a small tray with 1cm of water in the bottom underneath the cigarette to collect falling ash and minimize fire risk. Note: This step is not required if using e-cigarettes

5. Light the conventional cigarette and initiate the smoking protocol according to FTC guidelines, consisting of 35 ml draw, 2 seconds puff duration, 58 s intervals, and eight puffs per cigarette [7]. A full cycle consists of 2 cigarettes smoked back-to-back

6. For e-cigarettes, instead, the CORESTA vaping protocol should be applied. This consists of 55 mL draw, 2 puffs / 60 seconds, and 3 seconds puff duration. Note that the interval between puffs should be 27 seconds to keep the rate at 2 puffs / 60 seconds.

7. Following each cycle of exposure, remove mice from the gas induction chamber and return them to their cage with access to food and water for 30 minutes, and then repeat the procedure

8. Animals need to undergo through six to 8 cycles of exposure/day, every day for three weeks, although longer exposure times 4 to 8 weeks, can be used if needed.

Figure 1: Smoking machine setup.

Panel A: The smoking machine and the induction chambers housing the animals reside inside a chemical hood. Panel B: excerpts of the key setup components for in vivo experiments.

4. Experimental stroke: tMCAO

4.1. Materials

4.1.1. Animals

Mice groups that have completed the chronic smoke/vape exposure cycles plus gender-matched controls. Ideally, 6 to 8 animals/group are required for a basic study and allow for a margin of safety for unintended failed tMCAO procedures

Before initiating the tMCAO procedure, it is important to draw blood samples and perform a set of locomotor activity and exploratory behavior assessments (as described in section 4.5) to assess the pre-stroke baselines parameters, including glucose levels and any marker of interest. Measurements of blood concentrations of nicotine and cotinine are also important to evaluate the level and consistency of the exposure between animals (see Note 3)

4.2. Equipment:

1. Surgical Tools

   • 5–0 monofilament suture
   • Cauterizer
   • Heating pad
   • Rectal temperature probe
   • Electric clippers
   • Scalpel
   • Retractors
   • Curved scissors
   • Vannas-style spring scissors

2. Stereo dissection microscope

3. TR-200 homeothermic temperature system

4. Micrometer

5. Doppler flowmeter

4.3. Supplies

1. 70% ethanol

2. Isoflurane

3. Preservation materials

4. Brain Matrix Slicer

5. 2,3,5-triphenyl tetrazolium chloride (TTC)

6. PBS

7. Formalin

4.4. Methods

1. Sterilize all surgical tools by autoclaving (minimum 121 °C, 15 PSI, for 15 min). Sanitize the surgery table and associated equipment using 70% ethanol.

2. Cut a 5–0 monofilament suture (Harvard Apparatus, Holliston, MA) into 20 mm segments.

3. Round the tip of each segment by heating it near a cauterizer (Braintree Scientific, Inc., Braintree, MA). Measure the diameter of the tip using a micrometer (Applied Image Inc., Rochester, NY). We use a suture with a final tip diameter of 0.21–0.22 mm for a mouse with a bodyweight of 25–30 g.

4. Anesthetize the animal with 5% isoflurane (Aerrane, Baxter, Deerfield, IL) in 30% O₂ / 70% N₂O using the V-10 Anesthesia system (VetEquip, Inc., Pleasanton, CA). Following induction of anesthesia, reduce the level of isoflurane and maintain it at 1.5%.

5. Place the mouse in the supine position on a heating pad. Insert a rectal probe and monitor and maintain body temperature between 36.5–37.5 °C using the TR-200 homeothermic temperature system (Fine Science Tools Inc., Foster City, CA).

6. Shave the fur on the ventral neck region with electric clippers (Braintree Scientific) to expose the skin. Disinfect the surgical site using three applications of 70% ethanol.

7. Under a stereo dissecting microscope (Nikon, Japan), make a 1.5 cm long midline incision on the neck. Use retractors (Braintree Scientific) to expose the surgical field and identify the right common carotid artery (CCA), external carotid artery (ECA), and internal carotid artery (ICA). Carefully dissect the arteries free from surrounding nerves and fascia (see also Fig. 2).

8. The cerebral blood flow needs to be continuously monitored throughout the surgery to confirm the occlusion and reperfusion of the brain by using a non-invasive system such as a doppler flowmeter to evaluate the changes in microcirculation in the whole brain over different post-surgical periods.

9. Dissect the ECA further distally and coagulate the ECA and its superior thyroid artery (STA) branch using a bipolar coagulator (Howard Instrument Inc., Tuscaloosa, AL). Cut the ECA and STA at the coagulated segment.

10. Loosely tie two 8–0 silk sutures around the ECA stump. Apply a vascular clamp (Fine Science Tools) at the bifurcation of the CCA into the ECA and ICA.

11. Make a small incision at the end of ECA stump with Vannas-style spring scissors (Fine Science Tools). A 6–0 nylon monofilament with a rounded silicone-coated tip (0.20–0.23 mm; Doccol Corporation) is then gently introduced into the incision and advanced to the clamp. Tighten the two silk sutures around the lumen just enough to secure yet preserve the mobility of the in-dwelling monofilament suture.

12. Remove the clamp from the bifurcation. Gently advance the monofilament suture from the lumen of the ECA into the ICA for a distance of 9–10 mm beyond the bifurcation of CCA to occlude the origin of MCA. The duration of surgery is about 30–45 min.

13. Suture the incision on the neck and place the mouse in a 35 °C nursing box to recover from anesthesia and return it to the cage. It generally takes 5–10 min for the mice to recover from anesthesia.

14. The suture is then withdrawn up to the left carotid bifurcation to restore the blood flow (i.e., reperfusion; see Note 6). In the absence of other comorbid factors, the time interval between the onset of stroke (occlusion) and reperfusion (usually ranging from 30 minutes and 2 hours) determines the severity of the resulting brain damage.

15. Neurological evaluation using several sensory-motor tests (described below) are carried out after a preset time frame following reperfusion, depending on the study goals

Figure 2:

Schematic illustration of a basic MCAO procedure

4.5. Data acquisition and analysis

4.5.1. Behavioral tests:

Neurologic evaluation is carried out 96 h after reperfusion. The mouse will be held gently by the tail, suspended at about 50 cm above the bench, and monitored for forelimb flexion. Extension of both forelimbs straight toward the floor will be considered the absence of a neurologic deficit, and a score of 0 is assigned to the animal. A score of 1 (mild neurological deficit) is assigned to animals flexing the forelimb contralateral to the injured hemisphere. Mice will then be placed on a large sheet of soft plastic-coated paper to ensure that the animals can grip firmly with their claws. While holding from the tail, mice will be pushed behind the shoulder with gentle lateral pressure until the forelimb slide several inches. A score of 2 (moderate neurological deficit) will be assigned to animals showing decreased lateral push and forelimb flexion resistance. Lastly, a score of 3 (severe neurological deficit) will be assigned to animals showing the same behavior as for score 2, plus a circling movement.

4.5.2. Open-field tests:

Open-field tests evaluate the experimental animals’ locomotor activity and exploratory behavior. Animals are placed in the monitoring chamber to acclimate to the environment 20 – 30 minutes before the experiment and then monitored for 30 minutes using a SuperFlex Open Field system (Omnitech Electronics, Inc. Columbus, OH) or a similar apparatus. This system allows for tracking the location and certain behaviors of a subject animal via photosensors which create a 16×16 infrared grid. The test animal’s movement interferes with the infrared beams. The core software records and analyzes these interferences to correlate the motor behavior with tMCAO-induced neuronal damage (see also Note 7 and Fig. 3).

Figure 3: Open field test as a standard measure of exploratory behavior and general activity in rodents.

As shown in panel A, mice are housed into 16″ × 16″ unobstructed glass chambers containing infrared sensors along the perimeter. After the mice are introduced into the chamber (B), they are left to acclimatized to the new environment for about 30 minutes with no monitoring. Following the acclimatization, the monitoring and motion recording is initiated and continue uninterrupted for an additional 30 minutes. Automatic calculation of the activity (total distance traveled) and resting time of the animals is performed by Fusion software (C).

4.5.3. Terminal procedure:

Following the neurological assessments, anesthetize the mouse with 5% isoflurane and euthanize it by cervical dislocation. Decapitate the mouse and collect the brain. Slice the brain coronally into four 2-mm slices with a brain matrix (Braintree Scientific) on ice. Incubate the brain slices in 2% 2,3,5-triphenyl tetrazolium chloride (TTC) (Sigma-Aldrich) in 1X PBS for 20 min at room temperature to determine the size and extent of the infarction. Fix the brain slices in 10% neutral buffered formalin solution (Sigma-Aldrich) at 4 °C until imaging. The extent of infarction can be quantified as described elsewhere (see JoVE protocol 955)