A comparison of nicotine dose estimates in smokers between filter analysis, salivary cotinine, and urinary excretion of nicotine metabolites

Rationale Nicotine uptake during smoking was estimated by either analyzing the metabolites of nicotine in various body fluids or by analyzing filters from smoked cigarettes. However, no comparison of the filter analysis method with body fluid analysis methods has been published.
Introduction
A number of studies were published over the last 30 years that attempted to determine the amount of tar and/or nicotine that smokers receive from their cigarettes (reviewed in Stephen et al. 1989; Pritchard and Robinson 1996; Scherer 1999). The methodology used falls into three broad categories: (1) the analysis of biomarkers in human body fluids or expired breath; (2) the measurement of smoking behavior (puff volume, duration, and frequency) followed by a smoking machine set to duplicate human puffing conditions; and (3) the analysis of spent cigarette filters and the calculation of smoke yields from the filter efficiency.
Typically, biomarker measurements in blood/plasma and smoking behavior measurements require that sampling or measurements be made in a laboratory environment. There is a possibility that smoking behavior becomes atypical in this type of environment (Comer and Creighton 1978; Ossip-Klein et al. 1983). Urine, saliva, and spent cigarette filters can be collected in a smoker’s everyday environment. The analysis of 24-h urine samples for nicotine and major metabolites can provide quantitative data regarding uptake of smoke constituents as the product is used in a smoker’s everyday environment (see Byrd et al. 1998 and references therein for examples). Subject compliance can be an issue when trying to determine actual cigarette yields because in an unmonitored environment the subject must be relied upon to smoke only a given brand, not use any other nicotine-containing products, provide an exact accounting of every cigarette smoked during the collection period, and collect all urinary output.
The filter analysis method is one of the least invasive of the methods mentioned above. The smokers can use their product in their normal environment and the only deviation from normal behavior is to save the filters. Compliance is not an issue when trying to determine the subject’s cigarette yield because the filters from the actual cigarettes smoked are analyzed. In most cases, the returned filter can be compared to those of the subject’s stated brand to assure brand compliance. The primary issue with this method has been that smoking behavior can produce changes in the filtration efficiency of the filter. The filtration efficiency can vary according to the velocity of the smoke passing through the filter and, to some extent, the length of the tobacco rod smoked (Overton 1973; Dwyer and Abel 1986; Norman et al. 1984). The method used for this study was developed to minimize the effects of smoking behavior on filter efficiency by measuring only the portion of the filter downstream of the ventilation holes (i.e., the mouth end) (St.Charles 2001; Shepperd et al. 2006). This results in relatively constant filtration efficiency over a wide range of smoking behavior.
The objective of this study was to compare the nicotine yield of human-smoked cigarettes (mouth exposure) as measured by the filter analysis method and human smoke uptake as measured by biomonitoring under strictly controlled conditions. The biomonitoring measurements included salivary cotinine and 24-h urinary nicotine, cotinine, 3′-hydroxycotinine (3-HC), and their respective glucuronide conjugates. With a good correlation between the methods, future studies on smoker exposure can use the simpler filter analysis method rather than resorting to human biomonitoring techniques.
Materials and methods
Study design
The clinical portion of this study was conducted by an independent contract research organization in 2003 (Covance Clinical Research Unit, Madison, WI, USA). The analysis of salivary cotinine and urinary nicotine metabolites was performed at Covance Laboratories (Harrogate, North Yorkshire, UK). Filter analysis was performed by the study sponsor (Research and Development, Brown & Williamson Tobacco Company, Macon, GA, USA). The study was approved by Covance’s Institutional Review Board and performed in accordance with applicable federal regulations. Subjects who participated in the study gave their informed consent, were told of the purpose of the study, and could withdraw at any time.
Subject selection
Habitual smokers were recruited by Covance. Enrollment criteria included males or nonpregnant, nonlactating females, between 21 and 65 years of age, within −20% to +30% of their ideal body weight, who smoked at least 15 cigarettes a day of the same cigarette brand during the previous year. Subjects were excluded if they were under 21 years of age, were pregnant or lactating, participated in any other clinical study within 30 days before study entry, had a history or showed signs of a significant medical or psychiatric condition, used prescription medications within 14 days before study entry, had a history of alcoholism or drug addiction within a year of study entry, or used alcohol or any nonprescription preparations within 72 h of study entry. A few subjects deviated from the enrollment criteria: underweight (1), overweight (5), medication or alcohol usage before study entry (4), low cigarette consumption (2), shortened brand loyalty duration (4), elevated clinical chemistry (2), abdominal/hernia surgery (7), and positive drug screen before study entry (2). Because these deviations were considered minor and not expected to interfere with the study objectives, the subjects were allowed to participate in the study.
Subjects were assigned into one of four tar yield groups, which span the range of Federal Trade Commission (FTC) tar yields found in commercially available USA filtered cigarettes: 1–3 mg (ULL or ultralights/low), 5–6 mg (ULH or ultralights/high), 9–12 mg (LTS or lights), and 13–19 mg (FF or full flavor). The purpose was to cover a wide range of human nicotine exposure to allow robust correlations between the methodologies. The goal was to enroll 20 smokers per group; however, even with additional recruitment attempts, only 15 smokers enrolled in the ULL group (market share <2%). One subject in the FF group withdrew from the study due to illness. Table 1 summarizes the subjects’ demographics and their respective brand characteristics by tar band.
Table 1Subjects’ demographics and cigarette brand characteristics by tar bandSubjectsULLULHLTSFFTotalNumber1520201974Sex (M/F)8/712/810/1014/544/30Height (cm)173 (159–204)174 (157–194)174 (161–187)177 (165–190)175 (157–204)Weight (kg)77 (57–104)75 (42–98)78 (56–98)77 (60–100)77 (42–104)Age35 (21–54)32 (21–47)34 (21–64)32 (21–53)33 (21–64)Menthol smokers335718Brands Menthol/nonmenthol2/53/64/84/613/25 Tar (mg/cig)1.4 (0.7)5.4 (0.5)10.1 (0.9)15.0 (1.7)8.3 (5.1) Nicotine (mg/cig)0.15 (0.05)0.46 (0.04)0.77 (0.06)1.07 (0.14)0.64 (0.34) CO (mg/cig)1.9 (0.9)6.8 (0.6)11.6 (1.4)13.9 (2.0)8.9 (4.6) Puffs/cig7.6 (0.7)8.0 (1.1)7.9 (1.0)8.2 (0.9)7.9 (1.0)Height, weight, and age are shown as the mean (range). Cigarette yields are shown as the mean (SD) for the FTC smoking method.
Subjects were confined to the clinic for six calendar days to give five consecutive 24-h periods. Nine confinement periods were staggered and limited to ten subjects or less (generally of the same tar range group). During confinement period number 8, one LTS smoker and two FF smokers were allowed to participate with six ULL smokers to complete the LTS and FF cells. Subjects were fed a standardized bulk diet that excluded grilled, smoked, or barbecued food items. Consumption of water and other nonalcoholic beverages was unrestricted. Subjects refrained from strenuous exercise. During confinement, subjects smoked their usual cigarette brand ad libitum in a dedicated smoking room equipped with ventilation and air filtration. Use of any form of nicotine other than the subject’s declared own brand was prohibited. Cigarettes were purchased locally.
Urine and saliva collection and analysis
Twenty-four-hour urine samples were collected from each subject for five consecutive days. Collections started at approximately 0800 hours (first void excluded) and ended at approximately 0800 hours the following day (first void included). Urine was collected in 3-l amber plastic containers and kept refrigerated throughout the collection period. No chemical preservatives were used. After each 24-h sample collection, volume and pH measurements were recorded, a sample was taken for creatinine analysis, and 2 × 5-ml and 4 × 500-ml aliquots were taken and stored frozen at −70°C until shipped. Aliquots were shipped under dry ice to the analytical laboratory and stored at −70°C until analysis.
Urinary nicotine, nicotine-N-β-glucuronide, cotinine, cotinine-N-β-glucuronide, trans-3′-hydroxycotinine, and trans-3′-hydroxycotinine-O-β-glucuronide were analyzed using solid phase extraction (SPE) and liquid chromatography with tandem mass spectrometric detection using a method developed and validated at the analytical laboratory (Analytical Procedure Covance no. 2002-010). Nicotine-d3, cotinine-d3 (Sigma-Aldrich, Poole, UK), and trans-3′-hydroxycotinine-d3 (Toronto Research Chemicals, North York, ON, Canada) were used as internal standards. Nicotine, cotinine (Sigma-Aldrich), trans-3′-hydroxycotinine, and the three corresponding glucuronides (Toronto Research Chemicals) were used as reference standards. Samples were incubated for 18–22 h at 37°C with β-glucuronidase, which enzymatically deconjugated the aglycones from their respective glucuronides. The free and cleaved aglycones and their respective deuterated internal standards were then extracted and analyzed.
Saliva samples were collected in sterile Salivette tubes (Sarstedt, Newton, NC, USA) from each subject for five consecutive days at approximately 1830 hours. On day 4, two additional saliva samples were collected at approximately 0830 and 1330 hours. Clinical staff supervised and timed the process to assure compliance. Collected saliva samples were immediately stored at −20°C, shipped under dry ice to the analytical laboratory, and stored at −20°C until analysis. Salivary cotinine was analyzed by a method developed and validated at the analytical laboratory (Analytical Procedure Covance no. HB-02-061) based on a previously reported method (Bentley et al. 1999).
Filter collection and analysis
To control cigarette brand accessibility and document cigarette consumption, cigarettes were issued by clinical staff individually. The used filter had to be returned before a subject received their next cigarette. Used filters were processed under the supervision of clinical staff by having the subject remove any tobacco particles from the used filter before depositing it in an individually labeled glass container. Used filters were collected for five consecutive days with each day starting/ending at the same time as urine samples. The filters generated daily by each subject were shipped by overnight carrier at ambient temperature to the analytical laboratory. Immediately upon receipt, a 10.0-mm portion of the mouth end (tip) was cut from the filter using a specially designed jig, which kept the cut-length constant and kept a single edge razor blade perpendicular to the filter. Tip-to-tip length variation using the jig was within 3%. The tips were stored in labeled 30-ml glass jars with Teflon-lined lids at −20°C until analyzed.
Nicotine yields from the human-smoked cigarettes were estimated by analyzing the tips for nicotine. Five separate machine smoking regimes were used to provide calibration curves for the filter tips of each brand style tested. Cigarettes from the same batches that the subjects smoked were used for calibration smoking. The smoking regimes were chosen to give a wide range of cigarette yields of approximately equal spacing. Table 2 shows the calibration puffing conditions for each tar band. These machine puffing regimes proved to cover the spread of human smoking results for all but 7% of the subject-days tested (5 ULH, 1 LTS, and 6 FF out of 370 total). Six of the 12 involved extrapolating by less than 0.1 mg of nicotine/cigarette and the maximum extrapolation was from 2.3 to 2.6 mg/cigarette.
Table 2Calibration smoking conditions by tar bandULLULHLTSFF35/60/T + 3/O35/60/T + 3/O35/60/T + 3/O35/60/T + 3/O35/60/T + 3/B35/60/T + 3/B35/60/T + 3/B35/60/3 P/O70/60/T + 8/O35/60/4 P/O35/60/3 P/O35/60/6 P/O70/60/T + 8/B70/60/T + 8/B70/60/T + 3/O70/60/T + 3/O70/40/T + 3/B70/40/T + 3/B70/40/T + 3/B70/40/T + 3/OData are presented as puff volume (ml)/puff interval (s)/length smoked/ventilation holesO Open, B blockedT + # indicates overtipping plus distance (mm); # P indicates number of puffs.
Calibration equations were calculated using a linear regression of nicotine yield as a function of tip nicotine. These equations were then used to estimate human-smoked cigarette yield from the measured tip nicotine. This method was validated in a separate study using duplicated human puffing profiles (Shepperd et al. 2006). Aging tests by the study sponsor have shown that tip nicotine values were constant when whole filters were stored in glass jars at ambient temperature for up to 31 days. For this study, tips were cut from the whole filters within 5 days.
One to nine tips per sample were extracted using either 20 or 40 ml of methanol containing 0.038 mg/ml decanol internal standard. Tips were extracted for 40 min using a flatbed orbital shaker at 200 rpm. All available tips were divided into at least three separate batches, which were extracted and analyzed on different days to average analytical variation. For samples from days 2, 4, and 5, extract absorbance at 310 nm was also measured to audit the nicotine analysis using an independent method. Absorbance gives a measure of the tar deposited on the filter (Sloan and Curran 1981; Shepperd et al. 2006) and we have found that absorbance per tip correlates linearly with nicotine per tip. For days 1 and 3, replicate vials of the extract were stored at −20°C and subsequently analyzed for nicotine to audit the nicotine analysis further. The root mean square difference between the nicotine replicates was 5.5 μg/ml (7.3%). A backup set of replicate vials was kept for use when outlying data points were identified by either of the auditing methods. Samples were retested when they were outside of the 95% confidence interval of the absorbance per tip vs nicotine per tip correlation (days 2, 4, and 5) or when nicotine replicates differed by more than 20% (days 1 and 3).
The extract was analyzed for nicotine by gas chromatography using an Agilent 5890 Series II with a flame ionization detector and a J&W Scientific 30 m Megabore® 0.53 mm ID, DB-Wax (1.0 μm film) fused silica capillary column (Agilent, Palo Alto, CA, USA). The UV absorbance of the extract was measured using an Ocean Optics (Dunedin, FL, USA) PC2000 spectrometer equipped with a fiber optic dip probe with a 2-mm path length. This gave absorbance values within the linear range of less than 2 without further dilution. Absorbance was measured on the same day the tips were extracted because it was found that extract absorbance declines with even overnight storage.
Results
Measured urinary concentrations for each metabolite were multiplied by their respective daily urine volumes and converted based on molecular weights to yield recovery results in nicotine equivalents. The sum of the nicotine equivalents for nicotine, cotinine, and 3-HC were calculated for each subject per day to give total daily urinary nicotine equivalents (UNE) in milligrams per day. Nicotine yield per cigarette was calculated from the mean nicotine per tip values for each subject-day and the appropriate calibration equation for the brand smoked. This was multiplied by the number of cigarettes smoked by the subject that day to give nicotine yield in milligrams per day.
This study design allowed us to achieve a wide range of nicotine exposure suitable for the correlation of the three methods of nicotine uptake estimation. Nicotine yield from the human-smoked cigarettes (mouth exposure) as measured by filter analysis ranged from 3.7 to 67.1 mg/day, UNE ranged from 3.1 to 48.4 mg/day, and saliva cotinine ranged from 70 to 866 ng/ml. The mean (SD) proportions of urinary metabolites, including the glucuronides, were 22% (9%) for nicotine, 35% (6%) for cotinine, and 43% (13%) for 3-HC of the total UNE measured. When expressed as a percentage of nicotine entering the mouth, the proportions were 19% (9%) for nicotine, 31% (11%) for cotinine, 39% (17%) for 3-HC, and 89% (25%) for total nicotine equivalents.
Statistical correlations
Observed values from the three methodologies were correlated with each other. Figure 1 shows the three plots and correlations for all data points. Figure 1a is a graph of the UNE vs daily nicotine yield estimated from filter analysis; Fig. 1b shows UNE vs saliva cotinine; and Fig. 1c shows daily nicotine yield from filter analysis vs saliva cotinine. Linear regressions were significant for both the slope and intercept (p < 0.001) for all three correlations. The best correlation was obtained with UNE vs daily nicotine yield (Fig. 1a, R2 = 0.66). The slope indicates that 67% of the variation in nicotine mouth exposure calculated from filter analysis appeared as variation in the six urinary compounds measured. The standard error of the regression (SER) was 4.9 mg nicotine/day. Saliva cotinine did not correlate as well with either UNE (Fig. 1b, R2 = 0.49, SER = 5.9 mg/day) or nicotine yield (Fig. 1c, R2 = 0.45, SER = 7.4 mg/day). Because the amount of nicotine entering the mouth was greater than the sum of the UNE, the standard error of the nicotine yield would be greater than that of the UNE due to scaling even if the correlations were equivalent. To allow an equal comparison, the standard error of the percent of the difference between calculated (using the regression equation) and measured values was calculated for the three correlations. The standard error calculated in this manner was 32 and 42% for UNE vs nicotine yield and saliva cotinine, respectively, and 41% for nicotine yield vs saliva cotinine. Summary statistics for all correlations are in Table 3.
Fig. 1Correlations of the three methodologies (individual measurements)Table 3Summary of the statistics for linear correlationsCorrelation (y : x) Intercept (95% CI)Slope (95% CI)R2Standard error (%)Urine : filterIndividual4.3 (1.2)0.67 (0.05)0.664.9 (32)mg/day : mg/dayAverage2.7 (2.0)0.74 (0.08)0.833.2 (21)Urine : salivaIndividual5.8 (1.6)0.042 (0.004)0.495.9 (42)mg/day : ng/mlAverage5.4 (3.2)0.043 (0.009)0.555.2 (36)Filter : salivaIndividual6.9 (2.0)0.048 (0.006)0.457.4 (41)mg/day : ng/mlAverage5.6 (4.0)0.052 (0.011)0.546.5 (33)Filter : urine (creatinine normalized)Individual8.7 (1.9)10.8 (1.3)0.427.6 (41)mg/day : mg/day/mmol creatinineAverage7.1 (3.9)12.1 (2.7)0.526.7 (32)Urine : FTCmg/cig : mg/cigAverage0.49 (0.13)0.66 (0.19)0.410.27 (42)Filter : FTCmg/cig : mg/cigAverage0.52 (0.17)0.85 (0.24)0.410.35 (38)Saliva : FTCng/ml : mg/cigAverage211 (58)184 (81)0.22119 (46)Urine Urinary nicotine equivalents, Filter daily nicotine yield estimated from filter analysis, Saliva saliva cotinine concentration, FTC FTC nicotine yield, CI confidence interval
The means of the five daily data values per subject are shown in Fig. 2 using the same format as Fig. 1. Again, for all correlations, the slopes and intercepts were significant (p < 0.01). The correlation for UNE vs nicotine yield (Fig. 2a) improved significantly as shown in Table 3. The slope increased to 0.74 while the intercept moved closer to zero. The saliva cotinine correlations (Fig. 2b,c) also improved, but not as dramatically as the urinary nicotine. The correlation for UNE vs saliva cotinine (Fig. 2b) had an R2 value of 0.55 with a standard error of 5.3 mg/day (36%). The correlation for nicotine yield vs saliva cotinine (Fig. 2c) had an R2 value of 0.54 with a standard error of 6.5 mg/day (33%).
Fig. 2Correlations of the three methodologies (5-day average per subject)
Other findings
Although the primary purpose of this study was to determine the statistical correlations between the three methods of estimating nicotine uptake, data collected during the experiment allowed for additional observations.
Creatinine normalization Normalization by urinary creatinine is useful when only a single or partial daily urine collection is taken. To test the effect of creatinine normalization, observed daily UNE were divided by their respective millimole creatinine. A correlation was calculated with nicotine yield as the dependent variable and UNE/mmol creatinine as the independent variable. The results are summarized in Table 3. Both the intercept and slope are significant (p < 0.001) but normalization with creatinine clearly degrades the correlation compared to the correlation without normalization. The findings suggest that normalization with creatinine adds another factor of variability, which has a degrading effect on R2, and therefore indicates that the use of 24-h urine samples without normalization is expected to provide more accurate results than the analysis of partial (<24 h) urine samples with normalization. This is in agreement with Heavner et al. (2006), who showed mechanistically that creatinine normalization may be appropriate for some, but not all, of the urinary metabolites of nicotine and that other methods of normalization may be more appropriate for spot urine samples.
FTC smoke yields Because this was a confined clinical study, subjects would not necessarily be expected to behave as if they were in their normal environment. However, because a wide range of FTC cigarette yields were tested, it was considered worthwhile to test the correlation between the measured FTC yields and the three methods of nicotine estimation. Correlation results are summarized in Table 3 for the 5-day mean values for the three methods vs the FTC values on a per subject basis. All correlations gave significant slopes and intercepts (p < 0.001).
Differences between tar band groupings The results for mouth exposure to nicotine (from filter analysis), UNE, saliva cotinine, and cigarettes smoked per day were grouped by tar band and the mean, standard deviation, and one-way analysis of variance (ANOVA) were calculated on a per subject basis (Altman and Bland 1997). The results are shown in Table 4. Group means increased in rank order with increasing group yield except for saliva cotinine and cigarettes smoked per day. All measurements except cigarettes smoked per day gave a significant (p < 0.05) effect of group by ANOVA. For each measurement, results with different letter assignments are significantly different at the 95% confidence level (Fisher test). Normalizing urinary nicotine metabolites with creatinine resulted in an increase in p value for effect of group and no significant differences between the FF, LTS, and ULH groups. All three groups were significantly higher than the ULL group. This again shows that creatinine normalization degrades the discriminating power.
Table 4Mean ± SD for measurements grouped by tar band. ULLULHLTSFFANOVA pNo. of subjects15202019–Mouth exposure (mg nicotine/cigarette)0.63 ± 0.18a0.94 ± 0.29b1.19 ± 0.33c1.37 ± 0.58c<0.0005Mouth exposure (mg nicotine/day)14.7 ± 5.6a22.0 ± 7.1b24.4 ± 6.4bc28.3 ± 12.6c<0.0005Urinary nicotine (mg/cigarette)0.59 ± 0.19a0.84 ± 0.27b0.96 ± 0.21b1.20 ± 0.41c<0.0005Urinary nicotine (mg/day)13.6 ± 5.7a 19.3 ± 6.8b19.8 ± 5.9b24.5 ± 8.9c<0.0005Creatinine normalized urinary nicotine (mg/day/mmol)0.93 ± 0.58a1.31 ± 0.51ab1.43 ± 0.58b1.45 ± 0.52b0.040Saliva cotinine (ng/ml)216 ± 107a347 ± 119bc314 ± 88b411 ± 149bc<0.0005Cigarettes/day22.8 ± 4.7a23.4 ± 3.6a21.2 ± 4.5a20.8 ± 4.2a0.190Cigarettes/day (self-reported minus measured)1.6 ± 5.1−1.0 ± 8.0−0.9 ± 5.4−0.1 ± 5.9Results with different letters are significantly different at the 95% confidence level
Self-reported vs actual cigarette usage Table 4 also shows the difference in self-reported and measured cigarette usage. Self-reported cigarette usage was questioned and compliance with respect to truthfulness can be a concern (Byrd et al. 1998). In this study, subjects were asked to estimate their cigarette usage as part of the recruiting process. This gave a basis to compare with the measured usage during the study (overall mean = 22 cigarettes/day, SD = 4.2, range 15.8 to 32.0). The mean value for the (5-day average− self-reported) usage per subject was −0.2 cigarettes/day (SD = 5.9, range −19.4 to +11.7). Even when broken down to the smaller tar band groups of 15 to 20 subjects, the mean difference in measured and reported usage was less than 2 cigarettes/day.
Within-subject variation Because the trial took place over five consecutive days, within-subject, day-to-day variation was calculated for each of the measured variables. In addition, on day 4, saliva samples were taken at approximately 0830, 1330, and 1830 hours for measurement of saliva cotinine to estimate within-day variation. Results are summarized in Table 5 and are expressed as a pooled (root-mean-squared) coefficient of variation. The pooled day-to-day variations for each variable were similar, ranging from 15.2% for cigarettes smoked per day to 18.1% for UNE per day. The within-day variation of 8.1% for saliva cotinine was approximately half the day-to-day variation of 15.7%. When a single factor ANOVA was calculated using saliva cotinine for the three time periods sampled on day 4, the variation between time periods was insignificant (p = 0.58). However, this could have been overwhelmed by the subject-to-subject variation in nicotine uptake. This factor was removed by dividing the individual measurement by the daily average per subject for all three time periods. A single factor ANOVA was significant for time of day (p < 0.001) with the mean (95% confidence interval) saliva cotinine values being 1.008 (0.015), 0.96 (0.012), and 1.03 (0.013) times the average daily value per subject for the 0830, 1330, and 1830 hour samples, respectively. Even though the time of day had a statistically significant effect, the practical differences were small.
Table 5Within-subject variation expressed as a pooled coefficient of variation (CV)AnalyteTypePooled CV (range)%Filter (nicotine yield/day)Day-to-day16.7 (6–36)Urine (nicotine equiv./day)Day-to-day18.1 (4–54)Saliva cotinine concentrationDay-to-day15.7 (4–47)Saliva cotinine concentrationWithin-day8.1 (0.8–16)Cigarettes/dayDay-to-day15.2 (3–36)
Discussion
All three of the estimation methods correlated significantly with each other, but the best overall correlation was between the filter analysis method and UNE. The slopes for the correlations of UNE as a function of mouth exposure to nicotine were 0.67 and 0.74 for the individual and 5-day average regressions, respectively, implying that about 70% of the difference in the UNE measured in this study was due to a difference in mouth exposure to nicotine. However, the mean of the total UNE expressed as a percentage of the nicotine entering the mouth was 89%. The mean value falls within the reported range of 80% (Benowitz et al. 1994) to 90% (Curvall et al. 1991) but the slopes fall below this range. The difference between the two methods is due to the significant intercept calculated using the linear regression. The intercept using the 5-day average results was lower than the intercept using the individual results and the slope was greater. The intercept could represent compartmental nicotine and/or metabolite storage with subsequent carryover from storage before entering the study. This influence should diminish with a 5-day average compared to using single day results. An example of this is demonstrated in Fig. 1a where there is a single circled data point that appears to be an outlier. Urinary output of nicotine equivalents was approximately two times the mouth exposure of nicotine for that day. However, this was a day 1 measurement for a single subject. For subsequent days, the data points for this subject are buried within all the other data points. One possibility is that the subject had a much larger exposure to nicotine before participating in the study and the unusually high urinary output was due to clearance of the prior exposure. Averaging the results over 5 days would take out much of the metabolic influence and result in a much better correlation as demonstrated with these results.
For the saliva cotinine correlations, the R2 values only improved slightly with averaging. Part of the reason for the improvement with all correlations can be attributed to averaging out measurement variation. This should be similar for all correlations. However, the additional improvement in the filter vs urine correlation can be explained by metabolism. Nicotine has a relatively short serum elimination half-life of 2.2–2.9 h (Scherer et al. 1988; Benowitz and Jacob 1993; Benowitz et al. 1999, 2002, 2004), which means that the urinary nicotine should be from the nicotine taken in that day. Cotinine has a serum elimination half-life typically reported at 16–18 h (Scherer et al. 1988; Benowitz et al. 1999, 2002, 2004; De Schepper et al. 1987) and a similar urine elimination half-life (Benowitz and Jacob 1993; De Schepper et al. 1987). 3-HC, which is downstream metabolically from cotinine, has a serum elimination half-life of 5.9–6.6 h with a similar urine elimination half-life (Scherer et al. 1988; Benowitz and Jacob 2001). Given the pharmacokinetic information, these urinary metabolites must have originated from nicotine exposure on multiple days as has been demonstrated directly using nicotine infusion (Scherer et al. 1988).
Because measurements were taken over sequential days of input (filter analysis), the temporal source of urinary and salivary metabolites could be estimated by multiple regression of the metabolites vs the daily nicotine exposure. This analysis was performed using the nicotine exposure for the current day and the two previous days according to the equation:
where (N), (N-1), and (N-2) equals the nicotine exposure for the current day, 1 day before, and 2 days before the urine or saliva sample, respectively. Metabolite measurements from days 3, 4, and 5 were used. None of the coefficients for nicotine exposure from 2 days before the urine collection were significant (p > 0.15). For saliva cotinine, the coefficient for 2 days before was not significant at the 95% confidence but the p value of 0.07 was small enough to warrant consideration. Given this, the regressions were recalculated using (N) and (N-1) and including the urinary metabolite measurements from day 2 as well. The results of this regression are shown in Table 6.
Table 6Multiple regression results for source of urinary and salivary metabolites as a function of nicotine exposure from current and previous dayCoefficientSum of urinaryUrinary nicotineUrinary cotinineUrinary 3-HCSalivary cotinineR20.720.400.640.420.50Intercept3.51 (<0.001)0.77 (<0.01)0.85 (<0.01)1.89 (<0.001)97.9 (<0.01)Previous day0.24 (<0.001)−0.004 (0.89)0.08 (<0.01)0.16 (<0.001)5.7 (<0.01)Current day0.46 (<0.001)0.16 (<0.001)0.18 (<0.01)0.12 (<0.01)4.2 (<0.01)Results expressed as regression coefficient (p value)
For the sum of all urinary metabolites, the coefficients were significant for the current day and prior day’s exposure. This implies that the total urinary metabolites originated from nicotine exposure over 2 days. The only coefficient that was significant for the urinary nicotine regression was for the current day’s exposure as would be expected. The coefficient of 0.16 implies that 16% of the nicotine entering the mouth appeared as urinary nicotine plus the glucuronide. This agrees closely with the estimates of 13.8% (Benowitz et al. 1994) and 10–15% (Curvall et al. 1991). For the urinary cotinine regression, coefficients for the current day and previous day were significant with the current day being about twice that of the previous day. The two coefficients imply that 26% of the nicotine entering the mouth appears as urinary cotinine plus the glucuronide. This also agrees closely with estimates of 25.6% (Benowitz et al. 1994) and 20–25% (Curvall et al. 1991). For the 3-HC regression, the coefficients for the current day and previous day were significant with the current day being slightly smaller than the previous day. The sum of the coefficients (0.28) was smaller than the reported 41 to 60% of the total urinary metabolites (Benowitz et al. 1994; Curvall et al. 1991). It is possible that because 3-HC is the third step in the metabolism of nicotine, there was too much of a smoothing effect for the change in daily mouth exposure to fully capture.
For saliva cotinine, all coefficients were significant, suggesting that saliva cotinine results are an amalgam of at least 2 days of nicotine exposure. Saliva cotinine correlations suffer because results are expressed as concentration rather than an absolute value, and as such can be influenced by body size. In addition, other variables come into play that are not easily explained. An example of this is shown in Figs. 1 and 2 by the data points with boxes around them. These were from one subject that appeared to have unusually low saliva cotinine values for both UNE (Figs. 1b and 2b) and mouth exposure of nicotine (Figs. 1c and 2c). This subject had the highest mouth exposure of nicotine of all the subjects studied, yet the saliva cotinine values were only slightly above midrange. There was nothing unusual about this subject and average urinary output was within 1 SD (1,100 ml) of the average for all subjects (2,400 ml). It is unlikely that nicotine yields were overestimated because Fig. 1a shows that the data points are scattered about the regression line for UNE vs nicotine yield. It is also unlikely that the saliva cotinine concentrations were in error because the saliva cotinine for each day was analyzed in separate batches along with other samples. The subject did not rapidly metabolize cotinine to 3-HC because cotinine accounted for 47% of the urinary metabolites measured for this subject compared to an average of 35% for all subjects indicating the converse. Therefore, it must be concluded that this is an anomaly characteristic of this individual.
As shown in Table 3, with many of the estimation methods, the correlations are only slightly better than simply correlating with FTC nicotine yield of the cigarette. The average nicotine mouth exposure, as measured by the filter method, correlated with the FTC nicotine yield with a standard error of 38% and an R2 value of 0.41. The correlation of nicotine mouth exposure with saliva cotinine gave a standard error of 33% and creatinine-normalized urinary metabolites gave a standard error of 32% with R2 values approximately 0.1 higher.
Correlations of FTC nicotine with biomarkers in this study were stronger than that reported in other studies (Byrd et al. 1998; Jarvis et al. 2001; Ueda et al. 2002; Hecht et al. 2005; Bernert et al. 2005). We believe that there are valid reasons for this. One is that the correlations were performed using the 5-day averages per subject instead of a single sample per subject. In addition, exact compliance to brand and cigarette consumption was assured in the current study, whereas all but one (Bernert et al. 2005) of the referenced studies used self-reported brand identification and cigarette consumption. Self-reported brand information was reported to have about a 25% error rate when compared with packs returned (Peach et al. 1986) or in a test–retest comparison (Eisenhower et al. 1993). This and the potential for use of alternate brands during a study can further confound a correlation with biomarkers. Other confounding factors are the use of creatinine normalized spot urine samples instead of 24-h urine samples and analyzing for a subset of the nicotine metabolites used in the current study. Simple creatinine normalization is only biologically valid for xenobiotics that have the same excretion mechanism and urinary flow rate dependence as creatinine. Heavner et al. (2006) have shown that 1-HC (free and glucuronide) and cotinine glucuronide have urinary flow rate dependence similar to creatinine while nicotine (free and glucuronide), free cotinine, 1-hydroxypyrene, and the free and glucuronide forms of 4-(methylnitrosamino)-1-(3-pyridyl)-1-butanol do not.
In conclusion, two methods stand out as superior. One is the filter method, which estimates mouth level exposure directly on a per cigarette basis. Filter collection need not be quantitative, the filter returned can be compared to the brand it is supposed to be and it can be readily determined if it was smoked or not. Thus, brand compliance and smoking status can be assured even if the subject happened to occasionally use a different brand or other form of nicotine during the study. With self-reported daily cigarette use from groups of at least 15–20 subjects, the exposure per cigarette can be converted accurately to daily exposure. In addition, mouth exposure to tar can also be estimated using the filter method (Shepperd et al. 2006). The other measurement, which is considered by many to be the “gold standard,” is the measurement of urinary nicotine and metabolites from 24-h urine samples without creatinine normalization. This method appears to reflect the mean daily nicotine uptake of the last 2 days.

nicotine
filter
saliva
urine
metabolites
smoking behavior
cigarette smoking


Lack of evidence for direct phosphorylation of recombinantly expressed P2X2 and P2X3 receptors by protein kinase C

P2X3 and P2X2+3 receptors are present on sensory neurons, where they contribute not only to transient nociceptive responses, but also to hypersensitivity underlying pathological pain states elicited by nerve injuries. Increased signalling through P2X3 and P2X2+3 receptors may arise from an increased routing to the plasma membrane and/or gain of function of pre-existing receptors. An obvious effector mechanism for functional modulation is protein kinase C (PKC)-mediated phosphorylation, since all P2X family members share a conserved consensus sequence for PKC, TXR/K, within the intracellularly located N-terminal domain. Contradictory reports have been published regarding the exact role of this motif. In the present study, we confirm that site-directed elimination of the potential phosphor-acceptor threonine or the basic residue in the P+2 position of the TXR/K sequence accelerates desensitization of P2X2 receptors and abolishes P2X3 receptor function. Moreover, the PKC activator phorbol 12-myristate 13-acetate increased P2X3 (but not P2X2) receptor-mediated currents. Biochemically, however, we were unable to demonstrate by various experimental approaches a direct phosphorylation of wild-type P2X2 and P2X3 receptors expressed in both Xenopus laevis oocytes and HEK293 cells. In conclusion, our data support the view that the TXR/K motif plays an important role in P2X function and that phorbol 12-myristate 13-acetate is capable of modulating some P2X receptor subtypes. The underlying mechanism, however, is unlikely to involve direct PKC-mediated P2X receptor phosphorylation.
Introduction
P2X receptors constitute an abundant class of ligand-gated ion channels, which respond to extracellular ATP and related nucleotides with the opening of an intrinsic pore permeable to Na+, K+ and Ca2+ [1, 2]. A family encompassing seven P2X subunit genes, designated P2X1–7, has been identified in rodents and mammals. Six of the seven subunit isoforms (P2X1-P2X5 and P2X7) are able to assemble into homotrimeric receptor channels [3–8] with distinct pharmacological and electrophysiological phenotypes [9]. According to their sensitivity to the synthetic ATP analogue α,β-methylene adenosine 5′-triphosphate (αβ-meATP) and the rate of current desensitization, P2X receptors are generally subgrouped into at least two categories: (1) rapidly desensitizing and αβ-meATP sensitive (P2X1 and P2X3) and (2) slowly or non-desensitizing and αβ-meATP insensitive (P2X2, P2X4 and P2X7) receptors. The term “non-desensitizing” means that the currents are maintained for at least a few seconds in the continuous presence of agonist. A peculiar phenotype is inherent to heteromeric P2X2/P2X3 (P2X2+3) receptors, which feature αβ-meATP sensitivity combined with a non-desensitizing current response.
P2X receptors are found on the surface of a large variety of cells, where they are involved in numerous sensory processes including nociception under both physiological and pathological processes [10–15]. On nociceptive sensory neurons, extracellular ATP acts as a pain-producing neurotransmitter predominantly through homotrimeric P2X3 receptors or heterotrimeric P2X2+3 receptors. Conditions and mediators that facilitate P2X3 and P2X2+3 receptor-mediated signalling, apparently by potentiating ionic currents, include (1) substance P and bradykinin [16], (2) nerve injury [17] and (3) calcitonin gene-related peptide (CGRP), a potent vasodilator and proinflammatory agent [18]. How this increase in receptor function occurs has not been fully solved. One obvious candidate mechanism is protein kinase C (PKC)-mediated phosphorylation, since all P2X family members share a conserved consensus sequence for PKC-mediated phosphorylation (TXR/K) within the intracellularly located N-terminal domain (Fig. 1a, b). Indeed, results obtained with pharmacological activators and inhibitors are consistent with PKC being involved in P2X3 receptor potentiation by substance P and bradykinin [16], as well as the calcitonin gene-related peptide [18]. Direct PKC-mediated phosphorylation has so far been demonstrated only for the P2X2 receptor, with 18T of the 18TPK20 sequence as the phospho-acceptor site [19]. Mutational analysis has further shown that the TXR/K sequence of the P2X1, P2X2 and P2X3 receptors plays an important role in controlling the rate of receptor desensitization [16, 19, 20]. Changes of the rate of desensitization could profoundly influence the efficiency of synaptic transmission and thus contribute to pathological pain states. Overall, there is significant interest in a better understanding of the mechanisms involved in the short-term and long-term regulation of P2X receptors.
Fig. 1Effect of site-directed modification of a putative PKC motif on currents mediated by ATP-activated P2X2 and P2X3 receptors. a Schematic model of the transmembrane topology of the rat P2X3 subunit illustrating the N-terminal position of the 12TTK14 sequence. b Alignment of intracellular N-terminal amino acid sequences of the seven P2X subunit isoforms reveals a highly conserved consensus motif, TXR/K. c Typical current traces elicited by applying 10-s pulses of 100 μM ATP to oocytes expressing the indicated wild-type or mutant P2X2 receptors. d Typical current traces elicited by applying 10-s pulses of 100 μM ATP to oocytes expressing the indicated wild-type or mutant P2X3 receptors. Gray areas indicate the duration of ATP application. e All the P2X2 and P2X3 receptors and receptor mutants were expressed efficiently at the cell surface. Intact, healthy oocytes expressing the indicated proteins for 2 days were surface-labelled with the membrane impermeant reactive Cy5 dye and then extracted with dodecylmaltoside. Recombinant proteins were isolated by Ni2+ chelate chromatography and resolved by reducing SDS-PAGE. Shown is a fluorescence scan of an SDS-PAGE gel
The aim of this study was to assess biochemically the conditions in which P2X2 and P2X3 receptors become directly phosphorylated by PKC in a heterologous system. While this work was in progress, a similar study was published for the P2X3 receptor that arrived at essentially the same conclusion, namely that the P2X3 receptor is unlikely to be a direct PKC substrate [21]. Our results complement and extend this finding by demonstrating that the P2X2 receptor also does not serve as a substrate for direct PKC-mediated phosphorylation.
Materials and methods
Materials
The sources of antibodies, enzymes or peptide substrates are specified at the appropriate places in the text. Chemicals not otherwise specified were purchased in the highest available quality from Sigma-Aldrich (Taufkirchen, Germany) or Merck (Darmstadt, Germany).
Peptide-specific antibodies to P2X1, P2X2 and P2X3 subunits
Peptides ATSSTLGLQENMRTS (residues 385-399 of the P2X1 subunit), QQDSTSTDPKGLAQL (residues 458–472 of the P2X2 subunit) and VEKQSTDSGAYSIGH (residues 383–397 of the P2X3 subunit) were selected based on their lack of similarity with known proteins and predicted hydrophilicity for raising polyclonal antibodies against the C-terminal end of the rat subunits P2X1, P2X2 and P2X3, respectively. Rabbit polyclonal antibodies were ordered through the Custom Antibody Production Services of Eurogentec (Seraing, Belgium). After 3 months and a total of four immunizations, rabbits were sacrificed by complete bleeding. Peptide-specific antibodies were purified from the serum by affinity columns coupled with the peptides used for immunization. Immunoblotting showed that the antibodies recognized only the receptor against which they were raised, thus demonstrating isoform specificity of these antibodies (results not shown). The antibodies were aliquoted at 0.12–0.16 mg/ml in phosphate-buffered saline (PBS) containing 0.1% bovine serum albumin (BSA) and 0.01% thimerosal and stored at −80°C.
cDNA constructs
Plasmids encoding wild-type and N-terminal hexahistidine-tagged (His-tagged) versions of rat P2X1 subunits (rP2X1, GenBank accession no. X80477), P2X2 subunits (rP2X2, GenBank accession no. U14414) or rat P2X3 subunits (rP2X3, GenBank accession no. X90651) in the oocyte expression vector pNKS2 [22] were available from previous studies [3, 8]. Insertion and replacement mutations were introduced by QuikChange site-directed mutagenesis (Stratagene, La Jolla, CA, USA). For protein expression in mammalian cells, the receptor cDNAs were subcloned from the oocyte expression vector pNKS2 into pcDNA3.1. The plasmid His-rP2X2-EGFP-pcDNA3.1 with a C-terminal enhanced green fluorescent protein (EGFP) fusion was generated by eliminating the stop codon in His-rP2X2 and adding the coding sequence of EGFP (Clontech, Palo Alto, CA, USA) in frame. All constructs were verified by restriction analysis and nucleotide sequencing. A pEGFP-C1 plasmid harbouring the N-terminally EGFP-tagged splicing factor SF3B1 was kindly provided by our colleague Dr. Walter Becker [23].
P2X receptor expression in Xenopus laevis oocytes
Capped cRNAs were synthesized from plasmids linearized downstream to the 100 bp long polyA tail provided by the oocyte expression vector pNKS2 [22] and injected at 0.5 μg/μl in 50-nl aliquots into collagenase-defolliculated X. laevis oocytes using a Nanoliter 2000 injector (WPI, Sarasota, FL, USA), as detailed previously [24]. Oocytes were maintained at 19°C in sterile oocyte Ringer’s solution [ORi: 90 mM NaCl, 1 mM KCl, 1 mM CaCl2, 1 mM MgCl2 and 10 mM hydroxyethylpiperazine ethanesulfonic acid (HEPES), pH 7.4] supplemented with 50 μg/ml of gentamicin.
Heterologous expression in HEK293 cells
cDNAs encoding P2X subunits in pcDNA3.1 vector (Invitrogen, Karlsruhe, Germany) or the splicing factor SF3B1 in EGFP-C1 vector were transiently transfected into HEK293 cells using Lipofectamine LTX (Invitrogen, Karlsruhe, Germany). Cells were cultured at 37°C in Dulbecco’s modified Eagle’s medium (DMEM) (Invitrogen) supplemented with 10% (v/v) fetal calf serum (FCS) (PAA Laboratories, Linz, Austria), 100 U/ml of penicillin G and 100 μg/ml of streptomycin.
Metabolic [35S]methionine labelling of oocytes
For metabolic [35S]methionine labelling of oocyte-expressed recombinant proteins, cRNA-injected oocytes and non-injected controls were incubated overnight with L-[35S]methionine (>40 TBq/mmol; PerkinElmer, Boston, MA, USA) at about 100 MBq/ml (0.4 MBq per oocyte) in ORi at 19°C, and then chased for 24 h.
Surface fluorescence labelling of oocytes
Selective fluorescence labelling of plasma membrane-bound P2X receptors was achieved by using the amino-reactive fluorescent Cy5 dye (GE Healthcare, Freiburg, Germany), which is membrane impermeant due to its two sulfonic acid groups. Two days after cRNA injection, oocytes were washed with oocyte-PBS pH 8.5 (20 mM Na phosphate, 110 mM NaCl, 1 mM MgCl2), and then incubated for 30 min at ambient temperature with Cy5 dye, which was diluted 200-fold to a final concentration of 50 μg/ml from a dimethyl sulfoxide (DMSO) stock solution. The reaction was terminated by washing the cells with oocyte-PBS, followed by membrane protein extraction with dodecylmaltoside and receptor purification (see below).
Isolation of recombinant P2X receptors from X. laevis oocytes or HEK293 cells
His-tagged receptors were purified by Ni2+ nitrilotriacetic acid (NTA) agarose (Qiagen, Hilden, Germany) affinity chromatography from non-ionic detergent extracts of oocytes or HEK293 cells in the presence of iodoacetamide essentially as described [3, 8, 25]. Slight modifications were that buffers were supplemented with ethylenediaminetetraacetate (EDTA)-free Halt™ protease inhibitor cocktail (Pierce, Rockford, IL, USA), 50 mM NaF and 50 mM Na4P2O7 for phosphatase inhibition, and that digitonin was replaced by 0.2% and 0.05% dodecylmaltoside for membrane protein extraction and repetitive washing of resin-bound proteins, respectively. Proteins were released from the washed Ni2+ NTA agarose beads with non-denaturing elution buffer consisting of 250 mM imidazole/HCl (pH 7.4) and 0.02% dodecylmaltoside, and then kept at 0°C until analysed on the day of purification.
In addition, immunoprecipitation assays were performed using either cRNA-injected Xenopus oocytes or transiently transfected HEK293 cells. Cells were lysed in ice-cold immunoprecipitation lysis buffer containing 150 mM NaCl, 25 mM Tris/HCl pH 7.4, 20 mM NaF, 20 mM Na4P2O7, 10 μl/ml Halt™ protease inhibitor cocktail without EDTA (Pierce, Rockford, IL, USA), 1% sodium dodecyl sulfate (SDS) (w/v) and 50 mM iodoacetamide. For efficient solubilization, lysates were incubated for 30 min on ice with occasional vortexing, and then cleared by centrifugation for 15 min at 16,100 g and 4°C. The cleared SDS extract was diluted 1:1 with immunoprecipitation binding buffer [150 mM NaCl, 20 mM Tris/HCl pH 7.4, 0.4 mM phenylmethylsulfonyl fluoride (PMSF), 0.5% Triton X-100 v/v and 25 mM iodoacetamide], supplemented 1:1,000 with the desired peptide-specific P2X2 or P2X3 antibodies or a polyclonal goat anti-EGFP antibody (Rockland, Gilbertsville, PA, USA) and incubated for 2 h on ice. Immunocomplexes were bound to the protein A Sepharose by overnight head-over-head rotation at 8°C, then washed 5 times with immunoprecipitation binding buffer containing 0.1% Triton X-100 and finally eluted by incubating the beads for 15 min at 56°C in reducing sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer.
In vitro PKC phosphorylation assay
In vitro phosphorylation assays were performed by using P2X receptors as substrates, purified as detailed above. In addition, a selective peptide PKC substrate (RRGRTGRGRRGIYR, Calbiochem-Merck, Darmstadt, Germany, final concentration 150 μM) corresponding to the amino acid sequence 1487-1500 of the hepatitis C virus polyprotein [26] was used as a positive control. The proteins or the peptide were incubated for 30 min at 30°C with 0.02 unit (0.02 μg) of purified catalytic PKC subunit from rat brain and 50 μM [γ-32P]ATP (10 mCi/ml, Amersham Biosciences GE Healthcare, Freiburg, Germany) in phosphorylation buffer containing 6 mM CaCl2, 10 mM MgCl2, 20 mM Tris/HCl pH 7.5, 1 mM EDTA, 1 mM mercaptoethanol, 0.05% Triton X-100 (v/v) and 0.1 mg/ml BSA. The reaction was terminated by boiling in reducing SDS-PAGE sample buffer for 30 s. Proteins were resolved by reducing SDS-PAGE and visualized by immunoblotting or phosphorimaging as detailed below.
In vivo phosphorylation assay
Transiently transfected HEK293 cells were metabolically labelled by a 3-h incubation with 500 μCi/ml of [32P]orthophosphate (Amersham Biosciences GE Healthcare, Freiburg, Germany) in phosphate-free oocyte Ringer solution or in phosphate-free DMEM (Invitrogen, Karlsruhe, Germany) as appropriate. Cells were treated with or without 100 nM phorbol 12-myristate 13-acetate (PMA; Sigma-Aldrich, Taufkirchen, Germany) or 100 nM calyculin A (Tocris Bioscience, Ellisville, MO, USA) for 10 min at ambient temperature. Non-incorporated radioactivity was removed by rinsing the cells twice with tracer-free medium. Affinity purification or immunoprecipitation was performed as described above followed by SDS-PAGE and immunoblotting or phosphorimaging.
SDS-PAGE
Samples for SDS-PAGE were denatured by incubating in reducing SDS-PAGE sample buffer for 15 min at 56°C and then electrophoresed on a Laemmli SDS-PAGE gel (10% acrylamide) in parallel with [14C]-labelled molecular mass markers (Rainbow™, Amersham Biosciences GE Healthcare, Freiburg, Germany). Cy5-labelled proteins were visualized by scanning the wet SDS-PAGE gel with a fluorescence scanner (Typhoon, GE Healthcare, Freiburg, Germany). For visualization of radiolabelled or non-labelled proteins, SDS-PAGE gels were either fixed, dried and exposed to a PhosphorImager screen, and scanned using a Storm 820 PhosphorImager (Amersham Biosciences GE Healthcare, Freiburg, Germany) or/and blotted to polyvinylidene fluoride (PVDF) membranes.
Immunoblotting
Proteins separated by SDS-PAGE were electroblotted onto PVDF membranes. Membranes were blocked for 1 h with 5% non-fat milk in PBS (or, alternatively, with 5% BSA if the phosphothreonine monoclonal antibody was used for immunoblotting), washed 3 × 5 min with PBST (0.1% Tween 20 in PBS) and incubated for 2 h at 4°C in PBS containing the appropriate primary antibody. Phosphorylated proteins were detected by immunoblotting with a phosphothreonine monoclonal antibody (Cell Signaling Technology, Beverly, MA, USA). Alternatively, the peptide-specific antibodies to P2X2 and P2X3 subunits were used to verify receptor expression levels. Following washing in PBST, an appropriate horseradish peroxidase-conjugated secondary antibody (goat anti-mouse or goat anti-rabbit, Pierce, Rockford, IL, USA) was applied for 1 h in PBS supplemented with 1% non-fat milk. After another 3 × 5 min washes with PBST, immunoreactive bands were visualized by chemiluminescence imaging using a Fujifilm LAS-3000 system (Tokyo, Japan) and the AIDA Image Analyser Software Version 4.08 (Raytest, Straubenhardt, Germany).
Two-electrode voltage clamp current recordings and electrical capacitance measurements
Two-microelectrode voltage clamp measurements of X. laevis oocytes were performed 2 days after cRNA injection using a Turbo TEC-05 amplifier (npi electronics, Tamm, Germany) interfaced by an INT-20X AD/DA converter (npi electronics, Tamm, Germany) to a personal computer running the CellWorks Lite 5.1 software (npi electronics, Tamm, Germany), as detailed previously [27–29]. Current traces were elicited by 100 μM ATP at 1- to 5-min intervals (Sigma–Aldrich, Taufkirchen, Germany) at a holding potential of −60 mV in nominally Ca2+-free bath medium (designated Mg-ORi) to avoid activation of endogenous Ca2+-activated Cl− channels. ATP was prepared in Mg−ORi at its final concentration and applied by gravity-fed perfusion at a flow rate of 10 ml/min to an oocyte held by the two microelectrodes in the bath chamber (volume ∼10 μl). Switching between different perfusion solutions was achieved with electromagnetic valves controlled by the CellWorks Lite 5.1 software. Electrophysiological recordings were analysed using the Origin 6.0 software (Microcal Software Inc., Northampton, MA, USA). Single exponentials were fitted to the decay phase of the macroscopic currents with a nonlinear least squares method. The time constant τ was taken as an estimate of the rate of receptor desensitization.
To monitor PMA-induced changes in the oocyte surface area, depolarizing voltage steps of ΔV = 10 mV were applied from the holding potential to elicit a capacitive transient Icap, the integral of which yields the charge movement Qcap and thus the membrane capacitance Cm = Qcap/ΔV [30–32]. PMA was dissolved in DMSO to a stock solution of 10 mM and stored at −20°C. On the day of the experiment, the stock solution was pre-diluted to 100 μM in DMSO and then further diluted with Mg-ORi to the final concentration of 100 nM. Vehicle control experiments with a final concentration of 0.1% DMSO did not reveal any effects on P2X2 receptor-mediated currents measured with X. laevis oocytes (results not shown). All measurements were made at room temperature (21–23°C).
Data analysis
Data are shown as means±SEM. The paired Student’s t-test (two-tailed) was used to compare current amplitudes with statistical significance set to p < 0.05.
Results
Site-directed modification of a putative N-terminal PKC site affects the electrophysiological phenotype of P2X2 and P2X3 receptor channels
First, we tested each of the non-mutated and mutant receptor constructs for expression of an ATP-gated inward current in cRNA-injected X. laevis oocytes. Application of 100 μM ATP induced typical non-desensitizing currents through oocyte-expressed P2X2 receptors (Fig. 1c, left current trace). The P2X2 subunit shares with other P2X subunit isoforms a conserved consensus site for PKC phosphorylation (TXR/K, where T is the phosphorylation site and X denotes any amino acid; see Fig. 1a, b). Disruption of the conserved N-terminal PKC site by mutating the phosphor-acceptor 18T to A resulted in a rapidly desensitizing channel (Fig. 1c, middle current trace). Also, replacement of 20K by T resulted in a desensitizing phenotype (Fig. 1c, right current trace). Since PKC preferentially phosphorylates threonine (or serine) residues that are close to a C-terminal basic residue, replacement of 20K in the P+2 position by a non-basic residue such as T will destroy the putative PKC site despite preserving the phosphor-acceptor 18T. Virtually identical findings had previously been reported by Boue-Grabot et al. [19]. The average half-times of desensitization (τ) of the 18A and 20T P2X2 mutants were quite similar, with mean τ values (±SEM) of 4.5 ± 0.4 s and 7.3 ± 0.5 s, respectively, of eight oocytes from two females per value.
ATP applied to oocytes expressing the P2X3 receptor elicited a rapidly desensitizing inward current (Fig. 1d, left current trace) as is typical for this P2X receptor subtype. Incorporation of similar mutations, as in the P2X2 subunit in the analogous positions of the P2X3 receptor T12A or K14T, completely abolished P2X3 receptor function (Fig. 1d, middle and right current traces). Visualization of plasma membrane-bound fluorescence-labelled P2X receptors by affinity purification combined with SDS-PAGE and fluorescence scanning verified that all the receptors were efficiently exported to the oocyte surface (Fig. 1e).
The PKC activator PMA augments P2X3 receptor-mediated currents, yet leaves P2X2-mediated currents unchanged
Since the mutagenesis data for the P2X2 and P2X3 receptor were consistent with the possibility of a regulatory role for PKC in P2X receptor function, we examined whether the PKC activator PMA was capable of affecting the electrophysiological phenotype of P2X2 or P2X3 receptors. In planning the experiments, we considered that PMA treatment of X. laevis oocytes is known to induce a pronounced endocytotic reduction of the cell surface area, as evidenced microscopically and electrophysiologically by the disappearance of virtually all microvilli and a concomitant decrease of the electrical capacitance of the plasma membrane, respectively [31, 32]. Associated with oocyte surface reduction is a proportional internalization of membrane proteins, such as endogenous Na+-K+ pumps, suggesting that PMA treatment would stimulate internalization of recombinant P2X receptors as well. Re-examination of the time course of the PMA-induced reduction of the oocyte surface area showed that the electrical capacitance did not start to decline significantly before 10 min after PMA application (Fig. 2a). Accordingly, all current recordings were done within a time window of 10 min after PMA application to prevent any contribution of receptor internalization to the PMA effect on P2X receptor-dependent currents.
Fig. 2Effect of PMA on ATP-induced currents mediated by expressed P2X2 and P2X3 receptors in X. laevis oocytes. a The electrical membrane capacitance was monitored as a measure of the oocyte surface area during sustained stimulation with PMA. Each bar represents the mean Cm±SEM calculated from the areas under the current transients elicited in 10-s intervals by five consecutive 10-mV depolarizing steps. b, c Representative current traces elicited by 10-s lasting pulses of 100 μM ATP applied in 10-min intervals to oocytes expressing the indicated P2X receptors or P2X receptor mutants. Where denoted by a black bar, oocytes were pre-incubated with 100 nM PMA before ATP was co-applied. PMA was without effect on the P2X2 receptor-mediated current (b), but induced a marked increase of the current amplitude mediated by P2X3 receptors (c). Gray areas indicate the duration of ATP application
Figure 2b and c show typical current traces elicited by ATP from P2X2 or P2X3 receptor-expressing oocytes before and after a 10-min exposure to PMA. PMA neither affected the shape nor the amplitude of P2X2 receptor-mediated current (Fig. 2b). However, PMA produced a marked increase of the current amplitude mediated by the P2X3 receptor (Fig. 2c). In one experiment with four oocytes, a statistically significant increase of the P2X3-mediated current amplitude by PMA of 155 ± 42% (SEM) was determined.
P2X2 and P2X3 receptors exist in a non-phosphorylated state in both X. laevis oocytes and HEK293 cells
To explain the accelerated desensitization upon site-directed perturbation of the 18TXK20 motif, it has been suggested that the wild-type P2X2 receptor is constitutively phosphorylated at 18T and that this phosphorylation is responsible for the slow desensitization rate [19]. To address this issue, we next examined biochemically whether P2X3 receptors also exist in a constitutively phosphorylated state; the P2X2 receptor was also analysed to allow for a direct comparison of the results. Both parent P2X receptors and mutants were purified by Ni2+ chelate affinity chromatography from dodecylmaltoside extracts of [35S]methionine-labelled oocytes, resolved by SDS-PAGE, and blotted onto a PVDF membrane. However, probing with a phosphothreonine-specific monoclonal antibody revealed no discernible signal (Fig. 3a) despite the presence of significant amounts of [35S]methionine-labelled P2X2 and P2X3 protein in the SDS-PAGE gel (Fig. 3b).
Fig. 3Immunoblots show no constitutive phosphorylation of oocyte-expressed P2X2 or P2X3 receptors. The indicated wild-type or mutant P2X subunits were purified by Ni2+ chelate chromatography from [35S]methionine-labelled X. laevis oocytes and resolved by denaturing SDS-PAGE. a Shown is a representative immunoblot probed with a phosphothreonine-specific monoclonal antibody and a peroxidase-conjugated secondary antibody. No phosphorylation signal could be detected at the SDS-PAGE migration position of P2X2 and P2X3 subunits, which are indicated by the adjacent PhosphorImager scan in panel b. b Direct PhosphorImager visualization of metabolically [35S]methionine-labelled parent and mutant P2X subunits resolved by SDS-PAGE of the same samples as in panel a. cLeft panel: representative immunoblot of P2X2 subunits tagged with either an N-terminal or C-terminal hexahistidine sequence (designated His-P2X2 or P2X2-His, respectively) and probed with a phosphothreonine-specific monoclonal antibody and a peroxidase-conjugated secondary antibody. No phosphorylation signal could be detected at the SDS-PAGE migration position of P2X2 subunits, which were visualized by immunoblotting with the P2X2 subunit polyclonal antibody (middle panel) or PhosphorImager scanning of incorporated [35S]methionine (right panel)
To exclude the possibility that the apparent lack of receptor phosphorylation resulted from an interference between the N-terminal His tag and the PKC site of P2X3 or P2X2 receptor subunits, which are separated by only 12 or 20 residues, respectively, we performed analogous experiments with a C-terminally His-tagged P2X2 receptor, P2X2-His. As apparent from Fig. 3c, neither N-terminally nor C-terminally His-tagged P2X2 receptor became phosphorylated, as evidenced by immunoblotting with a phosphor-specific anti-threonine antibody (Fig. 3c). The presence of P2X2 receptor protein was verified by immunoblotting and metabolic labelling with [35S]methionine (Fig. 3c, middle and right panel).
Boue-Grabot et al. [19] have demonstrated phosphorylation of P2X2 receptors expressed in HEK293 cells. To address the possibility that P2X receptor phosphorylation is host cell specific, we analysed HEK293 cells transiently transfected with P2X2 or P2X3 expression plasmids. P2X receptor proteins were isolated by immunoprecipitation and resolved by reducing SDS-PAGE. The efficient expression of the P2X3 receptor or the C-terminally EGFP-tagged P2X2 receptor in HEK293 cells was verified by immunoblotting with subunit bands migrating at 54 kDa or 83 kDa (27 kDa contributed to the P2X2 subunit by the EGFP tag), respectively (Fig. 4b). However, phosphorylated P2X2 or P2X3 subunits could not be detected by immunoblotting with a phosphothreonine-specific antibody (Fig. 4a). In contrast, the PKC-phosphorylated splicing factor SF3B1, which was expressed in HEK293 cells as a positive control substrate and immunoprecipitated with an EGFP antibody, was efficiently phosphorylated under identical conditions, assuring that the assay was able to detect phosphorylated proteins (Fig. 4a, lane 1). Taken together, these results suggest that both P2X2 and P2X3 receptors exist constitutively in a non-phosphorylated state in both X. laevis oocytes and HEK293 cells.
Fig. 4Immunoblotting reveals no constitutive phosphorylation of HEK293 cell-expressed P2X2 and P2X3 receptors. The indicated proteins were isolated by immunoprecipitation using the peptide-specific P2X3 subunit polyclonal antibody or an EGFP-specific polyclonal antibody as appropriate and resolved by reducing SDS-PAGE. a Shown is an immunoblot probed by phosphothreonine-specific monoclonal antibody and a peroxidase-conjugated secondary antibody. Note that phosphorylation of the P2X3 receptor or the EGFP-tagged P2X2 receptor is not detectable, whereas a positive control—the EGFP-tagged splicing factor SF3B1 (migration mass 97 kDa [23])—was detected by the antibody. b The same samples as in panel a were immunoblotted using peptide-specific P2X2 and P2X3 subunit polyclonal antibodies to verify the expression of the transfected genes
PMA-activated PKC does not drive the phosphorylation of P2X2 or P2X3 receptors
A majority of protein kinases are quiescent unless their activity is stimulated by specific stimuli and effectors. To drive phosphorylation by active endogenous PKC, we incubated non-denatured recombinant His-tagged P2X1, P2X2 and P2X3 receptors isolated by Ni2+ NTA agarose affinity chromatography from dodecylmaltoside extracts of X. laevis oocytes with PMA-stimulated lysates of X. laevis oocytes (Fig. 5a) or HEK293 cells (Fig. 5b) in the presence of [γ-32P]ATP. As a positive control, a specific peptide PKC substrate RRGRTGRGRRGIYR was incubated with the lysates. This peptide was equally well phosphorylated by addition of either cell lysate or a commercial preparation of the rat brain PKC subunit (Fig. 5a, b). In contrast, however, no 32P incorporation could be detected in the SDS-PAGE migration position of the P2X1, P2X3 or P2X2 receptor subunit at 56 kDa, 54 kDa or 66 kDa, respectively. The presence of purified P2X1, P2X2 and P2X3 receptor subunit protein in this assay was verified by immunoblotting using P2X1-, P2X2- or P2X3-specific antibodies (Fig. 5c). The phosphorylation of the synthetic peptide clearly demonstrates that both X. laevis oocytes and HEK293 cells contain sufficient endogenous PKC to drive at least the phosphorylation of a synthetic peptide substrate.
Fig. 5Lysates of X. laevis oocytes or HEK293 cells contain endogenous PKC, but do not support PMA-driven phosphorylation of P2X2 or P2X3 receptors. The indicated wild-type and mutant P2X receptors were purified by non-denaturing Ni2+ chelate chromatography from X. laevis oocytes, supplemented with [γ-32P]ATP, 100 nM PMA and oocyte lysate (a) or HEK293 cell lysate (b), and incubated for 30 min at ambient temperature. In parallel, a peptide PKC substrate (final concentration 0.5 μg/μl) was incubated with [γ-32P]ATP, PMA and either of the two cell lysates or the purified rat brain PKC catalytic subunit as indicated. The proteins were resolved by reducing SDS-PAGE and visualized by phosphorimaging. Note that the band pattern was similar irrespective of whether protein of mock-injected oocytes or P2X receptor-expressing oocytes was used as a substrate, arguing against specific labelling of either P2X receptor. Arrows indicate migration positions of P2X1, P2X2 and P2X3 subunits. Arrowhead in a, dominant background band that is present in all oocyte samples. The peptide PKC substrate was similarly labelled by lysates or purified rat brain PKC, confirming the presence of endogenous PKC in X. laevis oocytes and HEK293 cells. c The same samples as in panels a and b were resolved on a separate SDS-PAGE gel. The indicated P2X subunits were detected by immunoblotting using the appropriate P2X subunit-specific polyclonal antibodies. A sample of mock-transfected cells was simultaneously probed with the P2X2 and P2X3 subunit-specific polyclonal antibodies
Discernible phosphorylation of P2X2 and P2X3 receptors might be obscured by a low level of endogenous PKC. Therefore, an analogous experiment was performed, in which the purified P2X2 and P2X3 receptors were incubated with purified rat brain PKC catalytic subunit rather than cell lysate in the presence of [γ-32P]ATP. Again, the specific peptide PKC substrate was efficiently phosphorylated as demonstrated both by direct phosphorimaging (Fig. 6a) and immunoblotting with the phosphothreonine monoclonal antibody (Fig. 6b). Phosphorylated P2X2 or P2X3 subunits could not be detected by either method (Fig. 6a, b) despite the presence of abundant amounts of the respective P2X subunits in the assays (Fig. 6c).
Fig. 6PKC phosphorylates a specific PKC substrate in vitro, but does not phosphorylate P2X2 or P2X3 receptors. The immunoprecipitated recombinant P2X2 or P2X3 receptor from X. laevis oocytes or a synthetic peptide (1 or 10 μg) derived from the non-structural protein 3 of hepatitis C virus, which serves as a specific substrate for PKC, were incubated for 30 min with purified rat brain PKC catalytic subunit (0.02 μg corresponding to 0.02 U/μl) in the presence of 50 μM [γ-32P]ATP (10 mCi/ml), and then subjected to SDS-PAGE and phosphorimaging. The rat brain PKC catalytic subunit does not require Ca2+ and phosphatidylserine for activity [42]. 32P was incorporated into the PKC substrate (left panel) and was also detected by immunoblotting with an anti-phosphothreonine antibody (middle panel). However, there was no detectable 32P incorporation at a migration position of ∼66 kDa or ∼54 kDa of the P2X2 or P2X3 subunit, respectively, the presence of which could be verified by immunoblotting (right panel)
PMA or calyculin A do not enhance phosphorylation of P2X3 receptors in intact HEK cells
In a final set of experiments, we attempted to directly demonstrate phosphorylation in vivo by using metabolic [32P]orthophosphate labelling combined with immunoprecipitation analysis as a read out. Mock-transfected and P2X3 receptor-transfected HEK293 cells were incubated with [32P]orthophosphate in phosphate-free DMEM, followed by incubation with the PKC activator PMA and/or the phosphoserine/phosphothreonine phosphatase inhibitor calyculin A. The PhosphorImager scan revealed an identical pattern of phosphorylated bands almost irrespective of the experimental conditions, with no evidence for a significant 32P-labelling of the P2X3 subunit (Fig. 7, top panel). P2X3 receptor expression was verified by immunoblotting (Fig. 7, bottom panel).
Fig. 7PMA and calyculin A do not induce phosphorylation of P2X3 receptors in intact HEK cells. Mock- and P2X3 receptor-transfected HEK293 cells were incubated with [32P]orthophosphate (500 μCi/ml) for 3 h in phosphate-free DMEM, followed by a 10-min incubation with 100 nM of each, the PKC activator PMA and/or the phosphatase inhibitor calyculin A. Triton X-100 extracts were then prepared from the cells and subjected to immunoprecipitation using the peptide-specific P2X3 subunit polyclonal antibody, followed by reducing SDS-PAGE and phosphorimaging. The migration position of the P2X3 subunit is marked by an arrow, indicating the absence of 32P incorporation into the P2X3 subunit. P2X3 receptor expression was verified by immunoblotting with the P2X3 subunit polyclonal antibody (bottom panel)
Discussion
Both P2X3 and P2X2+3 receptors are present on sensory neurons, where they play an important role in nociception. In addition to mediating transient nociceptive responses, these receptors contribute critically to the hyperexcitability that underlies abnormal pain states elicited by nerve injuries. One mechanism for the increased responsiveness to ATP is sensitization of P2X3 and P2X2+3 receptors arising from a significant enhancement in the trafficking of P2X3 subunit-containing receptors and/or an increase in function of pre-existing receptors [17]. An obvious candidate mechanism for functional modulation is phosphorylation, which is believed to occur universally in all ligand-gated ion channels as reviewed by Levitan [33].
Evidence for a control of P2X receptor channel function by PKC-mediated phosphorylation
Site-directed elimination of the putative N-terminal PKC site has already previously been shown to severely affect the electrophysiological phenotype of several P2X receptor subtypes, including P2X1, P2X2 and P2X3 receptors from rat and/or human [16, 19, 20, 34] and P2X5 receptors from frog [35]. Replacing the putative phospho-acceptor residue 18T or the basic residue in the P+2 position (20K) by A or a non-basic residue, respectively, imposed rapidly desensitizing properties onto the non-desensitizing wt P2X2 receptor without grossly affecting the current amplitude. In contrast, when analogous mutations were introduced into the rapidly desensitizing receptor subtypes P2X1 or P2X3, peak currents declined to low or undetectable levels, respectively [16 ,20, 34]; the same observation was also made with the slower desensitizing frog P2X5 receptor [35]. Overall, the results obtained by site-directed elimination of the N-terminal TXR/K motif can be plausibly reconciled with the hypothesis that the rate of P2X receptor channel desensitization is controlled by a phosphorylation-dephosphorylation mechanism, with phosphorylation at this site being associated with a decreased rate of desensitization.
Additional evidence for a possible involvement of PKC-dependent phosphorylation comes from experiments with the PKC activator PMA, which was consistently found to elicit a severalfold increase in currents mediated by the P2X1 receptor [36] or the P2X3 receptor [16, 18, 21] without changing the rate of desensitization. A PMA-induced change of the current desensitization was noted for C-terminally truncated P2X2 receptors, which are known to exhibit a desensitizing phenotype in contrast to the parent full-length P2X2 receptors [19, 37]. PMA treatment converted the rapidly desensitizing phenotype of truncated P2X2 receptors to slowly desensitizing, but did not affect the kinetic properties or current amplitudes of the full-length P2X2 receptors. The same lack of effect of PMA has been observed with wild-type P2X2 receptors stably expressed in HEK293 [38]. It has been inferred from these data that P2X2 receptors are constitutively phosphorylated and that this phosphorylation is responsible for their non-desensitizing phenotype. Consistent with this view, immunoblot analysis with a phosphothreonine-proline-specific monoclonal antibody efficiently detected wild-type P2X2 receptors with an intact PKC site, but not P2X2 receptor mutants with a disrupted PKC site [19]. Constitutive phosphorylation has been also demonstrated for the P2X1 receptor expressed in HEK293 cells [34]. Further studies demonstrated that exogenously added uridine triphosphate (UTP) potentiated the current mediated by endogenous and expressed P2X3 receptors in response to αβ-meATP as agonist. Substitution of phospho-acceptor residues of PKC consensus sites located in the ectodomain of the P2X3 receptor abolished this UTP-induced current potentiation, suggesting a role of ecto-protein kinase C in P2X3 receptor regulation [39, 40].
Evidence against a control of P2X receptor channel function by PKC-mediated phosphorylation
In direct contrast to the results of Boue-Grabot et al. [19], we were unable to demonstrate a direct phosphorylation of wild-type P2X2 receptors expressed in X. laevis oocytes and HEK293 cells. The various experimental approaches for phosphorylation detection that we applied included immunoblot analysis of purified P2X2 receptors. In the experiments shown, we used a phosphothreonine-specific monoclonal antibody instead of a phosphothreonine-proline-specific antibody, because the TXK motif of the P2X3 receptor contains a threonine in the -X-position instead of the proline found in the amino acid sequence of the P2X1 and P2X2 subunit (cf. Fig. 1b). However, even when we used the same phosphothreonine-proline-specific antibody as Boue-Grabot et al. [19], we could not demonstrate phosphorylation of P2X2 receptors in our experiments (data not shown).
Recently, it has been shown that a proline in the P+1 position strongly disfavours substrate recognition by PKC [41], suggesting that our negative results with P2X2 receptors may not result from technical limitations, but reflect the inherent inability of P2X2 receptors to serve as substrates for PKC. Notably, P2X3 receptors, despite harbouring a T in the P+1 position, could also not be demonstrated to become phosphorylated by PKC in our experiments, as well as in a recently published paper [21]. Further evidence against a direct PKC-mediated P2X receptor phosphorylation comes from co-expression studies of P2X1 and G protein-coupled receptors. Co-expression of the metabotropic glutamate receptor mGluR1α with P2X1 receptors produced a 2.5-fold increase of the P2X1 receptor-mediated current that was blocked by the PKC inhibitor staurosporine and could be mimicked by PMA treatment of singly expressed P2X1 receptors. Although the pharmacological data are basically consistent with PKC-mediated receptor phosphorylation being involved, mutant P2X1 receptors with a disrupted N-terminal PKC site (T18V) still showed PMA-mediated and mGluR1α-mediated potentiation. Also, PMA treatment did not lead to an increase in the (rather weak) basal phosphorylation of the P2X1 receptor [36].
Functional data supporting a role of PKC in P2X receptor regulation have been detailed above. There are, however, also functional data that are inconsistent with the view that the TXR/K motif represents a true PKC site: (1) replacement of the phospho-acceptor threonine by an acidic residue to mimic incorporation of a negatively charged phosphate group by constitutive phosphorylation was unable to restore the function of P2X1 receptors [34] and P2X3 receptors [16]; (2) the PMA-induced potentiation of hP2X1 receptor currents could be blocked by the PKC inhibitor staurosporine, but surprisingly, not by site-directed elimination of the PKC site [36]; thus it is much more likely that PMA exerts its effect on the P2X1 receptor (and the P2X3 receptor) through a yet to be identified accessory protein.
Taken together, there is no doubt that the TXR/K motif plays an important role in the functioning of P2X receptors, and that PMA is capable of modulating P2X1- and P2X3-mediated receptor currents. The underlying mechanism, however, is unlikely to involve direct PKC-mediated phosphorylation of P2X receptors. The existence of a proline in the P+1 position in several P2X subunit isoforms (P2X1, P2X2, P2X4) might be one causative factor preventing direct phosphorylation by PKC. Additional steric constraints may explain why P2X3 receptors also do not serve as PKC substrates, despite lacking the proline in P+1. The exact role of the TXR/K motif awaits further clarification.