Anal. Chem. 1987, 59, 437-439
437
Physiological pH Fiber-optic Chemical Sensor Based on Energy Transfer David M. J o r d a n and David R. Walt*
Max Tishler Laboratory for Organic Chemistry, Department of Chemistry, Tufts University, Medford, Massachusetts 02155 Fred P. Milanovich
Lawrence Livermore National Laboratory, University of California, Livermore, California 94550
A fiber-optic sensor has been developed contalnlng a fluore phore, eosin, and an absorber, phenol red, colmmoblilzed on the distal end of an optkai fiber. When an argon laser Is used to exclte eosin wRh light of X 488 nm, a reglon of the spectrum where phenol red does not absorb, eosin emlts light in a spectral regkn that overlaps signlfkantly wtth the absorpth spectrum of the basic form of phenol red. Consequently, nonradiative energy transfer occurs from eosin (donor) to phenol red (acceptor). The amount of energy transfer Increases as the pH increases resultlng in a dimlnlshed fluorescence intensity. Thus, changes in the absorption of phenol red as a function of pH are detected as changes in the fluorescent signal. I n thls manner a pH sensor optimized for physlologkai pH measurement has been prepared. The fiber exhibits a preclslon of at least 0.01 pH unlts.
Due to their small diameter and flexibility, fiber-optic chemical sensors offer an attractive possibility for invasive analysis of plant and animal tissues ( 1 , 2 ) . Blood gas sensors have been prepared for measurement of physiological p H (3-5), oxygen (6-9), and carbon dioxide (9). These fibers employ species-sensitive dyes that change their fluorescent properties in proportion to the concentration of the species of interest. The relatively low loss in attenuation of signals with distance in the optical fibers also permits applications such as remote monitoring of deep wells for groundwater analysis (IO). Fluorescence-based sensors have been employed in preference to absorption-based sensors because of increased sensitivity, greater latitude in geometric design, wide dynamic range, wavelength sensitivity, and linearity at low concentrations. Difficulties preventing practical applications of fluorescence spectroscopy to analysis have been described by Seitz (I) and by Crouch (11). Important among these are the “inner filter effects”, resulting from absorption of the excitation radiation by various species in the solution or matrix being analyzed (primary inner filter effect), or absorption of the emitted fluorescence radiation by these same competitive species (secondary inner filter effect). Correction schemes have been employed to compensate for these absorption effects but can be difficult to incorporate into fiber-optic technology. Another stringent requirement for developing fluorescence-based chemical sensors is the identification of an analyte-specific fluorophore. This latter restriction is severely limiting for most applications and is a key motive in developing absorbancebased systems. If a fluorescence-based method does not exist or is incompatible with an optical sensor, an absorption sensor is the only alternative. A multitude of absorbance-based analyses exist that would dramatically enhance the practical applications 0003-2700/87/0359-0437$01.50/0
of fiber-optic technology if the appropriate method for coupling this chemistry to a sensor could be developed. Only two absorbance-based fiber-optic sensors have been described (3, 12). These sensors employ scattering or reflecting methods for the absorbance measurement. As we had already developed a facile method for securing fluorescent measurements with fiber-optic technology (5), we decided to explore the possibility of linking an absorber’s specificity for a particular analyte to a fluorescent signal. We have now developed an absorbance-based, small-diameter, single-fiber chemical sensor with rapid response time, good stability, and high sensitivity that is useful for measuring physiological pH. This sensor is the result of a revolutionary design which should permit analysis of a wide variety of species. The sensor operates by the interaction of two moieties covalently attached to an optical fiber by copolymerization. One is a laser-stimulatable fluorescent dye which should be relatively insensitive to its environment; the other is a species, to be referred to as the “absorber”, which experiences changes in its absorbance properties proportional to the concentration of a specific analyte species. We envisioned that the presence of the absorber would modulate the fluorescence signal of the fluorophore. In principle, modulation of the fluorescence signal could be due to two distinct effects (13-15): 1. Inner Filter Effect. In this effect, absorbers cause diminution of fluorescence by absorbing either the excitation or emission radiation (vide supra). 2. Energy Transfer Effect. Upon excitation, the fluorescent molecule transfers its energy to an absorber nonradiatively. The absorber dissipates this energy through nonradiative processes. This effect is thus a fluorescence-quenching phenomenon. Both of these effects require that the fluorescence emission spectrum overlap with the absorption spectrum of the absorber (16). We chose to develop a pH sensor based on energy transfer from a pH-insensitive fluorophore, eosin, to a pH-sensitive absorber, phenol red. Phenol red was selected because, in the pH range 6.0-8.0, it absorbs in the same region that eosin fluoresces. Protonation of phenol red shifts its absorbance maximum to shorter wavelengths and thus causes diminished absorptivity in the overlap region. Therefore, as the pH decreases, the amount of energy transfer will be diminished, and eosin’s emission spectrum should show diminished quenching. Consequently, the efficiency of transfer should be dependent upon pH. EXPERIMENTAL SECTION Materials. Ammonium persulfate, N,N,N’,N’-tetramethyland riboflavin were from Bio-Rad ethylenediamine (TEMED), Laboratories, Richmond, CA; the (y-methacryloxypropy1)trimethoxysilane was obtained from Pharmacia, Piscataway, NJ. The 5-aminoeosin was obtained from Molecular Probes, Junction 0 1987 American Chemical Society
438
ANALYTICAL CHEMISTRY, VOL. 59, NO. 3, FEBRUARY 1, 1987
City, OR, or synthesized by bromination of 5-aminofluorescein in acetic acid solution. All other chemicals were obtained from Aldrich Chemical Co. All purchased reagents were used without further purification, except for tetrahydrofuran (THF), which was freshly distilled from lithium aluminum hydride in a nitrogen atmosphere just prior to use. Glass-on-glass fibers (200/250 pm), of length approximately 1m, were terminated at one end with amp connectors. The other ends, from which a length of approximately 2 cm of buffer was removed, were washed in concentrated sulfuric acid and rinsed with distilled water. The connection ends were polished and the stripped ends cleaved squarely as verified by microscopic examination. Glass capillary tubes were slipped over the fiber tips for protection and for aiding alignment of the fibers in the polymerization reaction chamber. Preparation of the pH Probe. Surface silanization of the fiber tip was performed by submerging the cleaned fiber tip into a solution of (y-methacryloxypropy1)trimethoxysilane(2% v/v) in water (acidified to pH 3.5 with acetic acid) for 1 h at room temperature. After silanization, the fiber was washed with pH 3.5 water and then used in the next step within 1 h. Aqueous solutions of acrylamide (caution: toxic) and N",methylenebis(acry1amide) (BIS) were prepared according to the procedure of Updike and Hicks (17). The N-(5-eosinyl)acrylamide was prepared immediately prior to use by mixing freshly distilled THF (1.0 mL) with 5-aminoeosin (0.043 g/0.06 mmol) and then adding acryloyl chloride (23 pL, 0.3 mmol) after which the mixture was allowed to stand in the dark at room temperature for 1 h. A polymerization medium was prepared by mixing 5 mL of acrylamide solution, 20 mL of BIS solution,and phenol red (0.125 g, 0.37 mmol), which was deoxygenated with nitrogen and maintained in a nitrogen atmosphere in a reaction chamber as previously described (5). To the reaction was added with stirring the THF solution of N-(5-eosinyl)acrylamide, ammonium persulfate (0.655 g, 2.9 mmol), and TEMED (50 wL, 0.33 mmol). The lid of the chamber, in which prepared fibers were positioned, was put in place. The mixture was allowed to stand at room temperature in a nitrogen atmosphere for an hour. If gelation had not occurred within this time, heating (to approximately 50 "C) brought about polymerization. When polymerization had occurred to the extent that the mixture's surface did not flow when the reaction chamber was tilted,the fibers were withdrawn from the reaction chamber. Clots of polymer not near the tips of the fiber were removed from the sides and the fibers were soaked overnight in a buffer of pH 7.4-7.9. Laser Apparatus. The instrumental configuration employed for excitation of the distal end of the fiber and for monitoring the fluorescence response has been described previously (5). An argon-ion laser, Spectra-Physics Model 162A-04,Mountain View, CA, provided the excitation radiation (typically 488 or 514.5 nm), which was passed through a neutral density filter and a dichroic mirror to the coupled fiber. The returning fluorescent signal (red shifted) returns via the same fiber, and is deflected 90" by the front surface of the angled dichroic mirror. This signal beam is long pass filtered, focused, and passed through a slit into a single grating monochromator. The resulting wavelength dispersed signal is measured with a photon-counting detection system (Pacific Instruments Model 126, Concord, CA). The intensity of the fluorescence is measured in photon counts per second as a function of time or of the wavelength examined. Measurements. The fiber, the distal end of which is extended from the glass capillary sheath, is positioned vertically. These solutions are raised to the tip by a labjack, taking care that the end of the fiber does not come into close contact with a reflecting surface such as the bottom of the glass container. Slight agitation of the solution prior to measurement is performed to ensure that residues of the previous sample are flushed from the region of the fiber tip. Laser excitation of the eosin has been performed at 488 and at 514.5 nm, the latter being near the absorption maxima of eosin. Power levels of 3.0 pW are routinely used, but fiber sensitivity to pH variation has also been demonstrated at power levels as low as 0.3 pW. The intensity of the fluorescence signal of the fiber is observed at 546-547 nm, the emission maxima. Buffer solutions were prepared in the manner of McIlvaine, and the pH values were verified by measurement prior to use.
o^ 0
z
200-
X VI
P v
.-b
--
g
100-
0
L 4
5
6
7
8
9
PH Figure 1. pH response of phenol red-eosin fiber. All measurements were taken at 546 nm with 2-nrn bandwidth and 488-nm excltation.
All pH response values were determined in replicate by cycling through the pH range, in both orderly and random fashion. The response time of the photometer was set to 0.3 s and intensity measurement data were obtained only after a lapse of three time constants for each buffer change. All intensity measurements were performed at room temperature. Scanning electron micrographs were performed on an AMRay 1000 B scanning electron microscope. Fibers were mounted in a thick coating of silver paste, air-dried overnight, and coated with 5 nm of Au/Pd (4060) with an Anatech, Ltd., Hummer X. Scanning angles were separately optimized for each fiber.
RESULTS AND DISCUSSION Figure 1displays the pH sensitivity of a fiber exposed to 15 buffers, in the pH range 4.49-8.60. The exciting laser beam was closed during buffer changes. The data show that the fluorescence intensity decreases as the pH increases. This result can be explained by either energy transfer or secondary inner filter effects. The decrease in intensity from pH 4 to pH 8 is 65%. Assuming a random distribution of phenol red in the polymer layer furnishes a concentration of 0.014 M. If we further assume an average path length of 3.5 pm (1/2 X 7 pm layer thickness, vide infra), we calculate that the secondary inner filter effect would account for -10% loss in signal. Therefore the predominant effect observed in this sensor is energy transfer. As the pH increases the spectral overlap integral between the eosin emission and phenol red absorption spectra increases. This increased overlap results in greater energy transfer from eosin to phenol red (16). The greater the transfer, the less fluorescence is detected. In this manner the fluorescence signal is modulated by the absorber which in turn is modulated by pH. The precision is A0.008 pH units and is based on the photoelectric detection of signal with the minimum detectable change being twice the noise of the measurement. The data in Figure 2 were obtained in an evaluation of the sensor's response time. During time period a, the shutter is opened to provide excitation to the sensor which has equilibrated in a buffer of pH 7.1. During period b, the intensity of fluorescence in this buffer is recorded. At the beginning of period c, a large quantity of pH 6.5 buffer is added. At the end of period c the sensor has had a 100% response, having established a new stable intensity level. If the definition of response time is the time required for 63% of maximal response (Peterson on pH), the response time of this fiber is 0.07 min. This is one-tenth the response time of the Peterson sensor (3) and is comparable to that of our previous fluorescence-based pH sensor. The 5-aminoeosin was readily derivatized with an alkene function by reaction with acryloyl chloride. Peterson e t al. have reported that phenol red may be incorporated directly into acrylamide polymers without derivatization ( 3 ) . This property allowed facile incorporation of phenol red into the
ANALYTICAL CHEMISTRY, VOL. 59, NO, 3, FEBRUARY 1. 1987
d
T i m e (min)
Flgure 2. Evaluation of fiber’s response thne: (a) shutter opened wlth fiber immersed In pH 7.1 buffer; (b) equilibrated s’ylnal in pH 7.1 buffer: (c) large quantity of pH 6.5 buffer added; (d) equilibrated signal for pH 6.5 buffer.
a
439
T o determine the sensitivity of these fibers to photobleaching, the sensors were exposed to continuous exciting radiation of 3.0 pW power. The signal (cps) did not change over a 10-min period in pH 4.5 buffer; however in pH 6.9 and 7.7 buffers, the signal increases were 1.4% and 6.0% respectively. The fact that fluorescence intensity increases rather than decreases, as is the case for fluorescein sensors, suggests that the phenol red component is photohleaching. Bleaching does not, however, seem to be a significant problem when making measurements by opening the shutter for short time periods; the intensity signals remain reproducible after cycling several times through a series of 10 to 12 buffers. The merits and methodology of preparing chemical sensors by the covalent attachment of dye-containing polymers to a glass fiber have heen previously described (5).This new sensor differs from the previous one in that it contains two immobilized dyes, and the concept of operation is based on modulation of fluorescence emission by reaction of a species other than the fluorescent dye itself. The phenol red-eosin prohe described in this paper has proven to he highly sensitive for reversible pH measurements in the physiological range. The probe is simple to prepare, is resistant to photobleaching, and is durable. We believe that further application of the concept demonstrated in the preparation of these energy transfer has& sensors will provide expanded capabilities for fiber optic sensor analysis. The me of optical fibers with thin polymeric coatings containing a mixture of a fluorophore and a species-selective absorber can lead to the preparation of specific small diameter sensors with rapid response times.
ACKNOWLEDGMENT The authors express their gratitude to Darrel Garvis and Christiane Munkholm for technical assistance and to Sylvie Lalihert6 of the USDA Human Nutrition Research Center at Tufts University for performing the scanning electron microscope studies. Registry No. H,C==CHCOCI, 814-68-6; eosin, 17372-87-1; phenol red, 143-74-8;(y-methacryloxypropyI)trimethoxysilane, 2530-85-0; 5-aminoeosin, 75900-75-3;N-(5-eosinyl)acrylamide, 105502-82-7; N-(5-eosinyl)acrylamide-N,”-methylenehisacrylamide-phenol red copolymer, 105502-83-8. LITERATURE CITED
Flgwe 3. Scanning electron micrographs: (a) bare liber, small dust and dirt particles can be seen on all surfaces: (b) phenol red-eosin fiber, peeling can be observed on the sides of the fiber indicating a thickness of approximately 1 0 sm.
polymer without further derivatization. We presume that t h i ~ copolymerization reaction of phenol red with acrylamide succeeds because it possesses quinonelike character. Scanning electron micrographs (Figure 3) show that the coating is uniformly thin with a thickness of approximately 10 pm. The thickness is determined by observing vacuuminduced peeling in various SEMs resulting from the fiber processing. The thinness and porosity of the polyacrylamide layer result in very short response times and 100% equilibration times of 10 s or less. Furthermore, the fiber can he repeatdy dried and rewet without loss of signal or sensitivity.
(1) Seitr. W. R. Anal. Chem. 1984, 56. 16A-34A. (2) Peterson. J. I.: Vurek. G. G. Scklnca (Washington. D.C.) 1984. 224 (13 April). 123-127. (3) Pe1erson. J. I.: Goldstein. S. R.: Fitrgerald. R. V. A w l . Chem. 1980. 5 2 . 864-669. (4) Ssari. L. A,: Sew. W. R. Anal. Chem. 1982. 54. 821-823. (5) Munkhdm. C.: Wan. D. R.: Milanovlch, F. P.: Klalner. S. M. Anal. Chem. 1988. 5 8 . 1427-1430. (6) Wolfbeis, 0.S.: Posh. H. E.: Kroneis. ti. W. A w l . Chem. 1985. 5 7 . 2556-2561. (7) Peterson. J. I.: Fitrgerald, R. V.: Buckhold. D. K. Anal. Chem. 1984. 56. 62-67. (6) Zhujun. 2.;Seitr. W. R. Anal. Chem. 1988. 58. 220-222. (9) Gehrich. J. L.: Lubbers. D. W.: Opitr. N.: Hanrmann. D. R.: Miller. W. W.: Tusa. J. K.: Valusi. M. IEEE Trans. Blamed. Eng. 1988. EM€-33. 117-132. (10) Milanauich. F. P.: Gawls. D.: Angel. S.M.: Klainer. S.:E C C ~ SL.. Anal. Insfrum. (N.Y.) 1986. 15. 137-146. (11) Ralriall. E. H.: Harlmnn. R. 0.:Crouch. S. R Anal. Chem. 1984. 56. 342-347. (12) Coleman. J. T.: Eastham. J. F.: Sepaniak. M. J. A m i . Chem. 1984. 5 6 , 2246-2249. Faraday SCC. 1959. 2 7 . 7-17. (13) FMSter. T. DISCUPS. (14) Kundy. P. C. Natwwissenschenen 1981. 4 8 . 644. (15) White. C. E. I n F I ~ s c e n c e - T M t y .Inshwnenlathm andPrac(lce: Guiibaun. G. G.. Ed.: Choler 7. Marcel Dekker: New Yark. 1967: _..“r.”.
..
(16) Haugland. R. P.: Yguerablde. J.: Stryer. L. Proc. Nan. ACdd. Sci. U . S . A . 1969, 23-30. (17) Hicks. G. P.: Updike. S. J. Anal. Chem. 1986. 38. 726-730.
RECEIVED for review August 14,1986. Accepted September 30, 1986. This work was supported by the Environmental Protection Agency through Tufts Center for Environmental Management.
