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Rapid Acid-Base Titrations Using Microfluidic Paper-Based Analytical Devices Shingo Karita, and Takashi Kaneta Anal. Chem., Just Accepted Manuscript • DOI: 10.1021/ac5039384 • Publication Date (Web): 25 Nov 2014 Downloaded from http://pubs.acs.org on December 1, 2014

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Analytical Chemistry

Rapid Acid-Base Titrations Using Microfluidic Paper-Based Analytical Devices Shingo Karita and Takashi Kaneta* Department of Chemistry, Graduate School of Natural Science and Technology, Okayama University, Okayama, 700-8530 Japan

1

ACS Paragon Plus Environment


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ABSTRACT: Rapid and simple acid-base titration was accomplished using a novel microfluidic paper-based analytical device (µPAD).

The µPAD was fabricated by wax

printing and consisted of ten reservoirs for reaction and detection.

The reaction

reservoirs contained various amounts of a primary standard substance, potassium hydrogen phthalate (KHPth), whereas a constant amount of phenolphthalein was added to all the detection reservoirs.

A sample solution containing NaOH was dropped onto the center of

the µPAD, and was allowed to spread to the reaction reservoirs where the KHPth neutralized it.

When the amount of NaOH exceeded that of the KHPth in the reaction

reservoirs, unneutralized hydroxide ion penetrated the detection reservoirs, resulting in a color reaction from the phenolphthalein.

Therefore, the number of the detection

reservoirs with no color change determined the concentration of the NaOH in the sample solution.

The titration was completed within 1 min by visually determining the end point,

which required neither instrumentation nor software.

The volumes of the KHPth and

phenolphthalein solutions added to the corresponding reservoirs were optimized to obtain reproducible and accurate results for the concentration of NaOH.

The µPADs determined

the concentration of NaOH at orders of magnitude ranging from 0.01 to 1 M.

An acid

sample, HCl, was also determined using Na2CO3 as a primary standard substance instead of KHPth.

Furthermore, the µ-PAD was applicable to the titrations of nitric acid, sulfuric

acid, acetic acid, and ammonia solutions.

The µPADs were stable for more than 1 month 2


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when stored in darkness at room temperature, although this was reduced to only five days under daylight conditions.

The analysis of acidic hot spring water was also demonstrated

in the field using the µPAD and the results agreed well with those obtained by classic acid-base titration.

3



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Recent advances in micro- and nano-scale analytical devices have resulted in excellent performance, which cannot be achieved by classic techniques.

For example, the

miniaturization of analytical devices has permitted a high degree of sensitive detection,1 rapid separation,2 and enhancement in the rate of chemical reactions.3

In particular,

microfluidic devices consisting of glass and polymer substrates are widely employed for miniaturizing several analytical devices including chemical sensors, separation channels, and manipulation channels of micro-sized particles and biological cells.4 Conversely, microfluidic paper-based analytical devices (µPADs) have recently attracted a great deal of interest due to a simple structure, easy fabrication, lightness of weight, inexpensive materials, and rapid analysis.

The first µPAD used a

photolithographic method to determine blood constituents such as protein and glucose.5 Thereafter, several improvements in the fabrication of µPADs have been explored using wax printing,6,7 ink-jet printing,8 screen printing,9 laminate film,10,11 and movable-type printing.12

Among these, wax printing and ink-jet printing are the simplest and most

precise ways to fabricate microfluidic channels on paper since conventional drawing software can be employed for designing µPADs. Using µPADs, some detection techniques have been applied for the determination of analytes, such as colorimetry for metal ions,13-16 biomarkers,17 DNA,18 and foodborne pathogens.19

Other techniques include enzyme assays20 and electrochemistry.21-24 4


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Colorimetry is perhaps the most popular since only a scanner (or a digital camera) and software for image processing are necessary for quantitative analysis, i. e., both the devices as well as the detection system are small, inexpensive, and easy to use.

A limitation of

µPADs that has been reported in past publications is the need to calibrate using standard solutions to quantify an analyte.

The intensity of the color for the sample must be

compared with those of standard solutions, and calibration curves must be constructed for quantitative analysis using a scanner and image processing software.

Only a few

exceptions have been demonstrated without the use of either a scanner or a digital camera. Cate and coworkers used the distance of color development to quantify analytes,25 and Lewis and coworkers employed time as a quantitative readout.26,27

In the µPAD based on

the distance of color development, as flowing analyte forms precipitation by the reaction with reagents, color develops along the flow channel until all of the analyte is consumed. When using the µPAD, however, the distances in color development still must be calibrated to the concentrations of the analyte using the standard solutions. Conversely, compared to modern instrumental analysis, classic analytical methods such as titrimetry and gravimetry need no calibration curve since the analyses are based on the absolute amounts of titrant and the weight of the precipitation.

For example, in acid-base

titration, if we use a primary standard solution as a titrant, or titrand, the concentration of an acid or a base solution is directly calculated from the volume required to reach the end 5



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point.

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In addition, titration permits on-site measurement using no instrumentation since

the human eye determines the end point.

However, this type of analysis requires a large

amount of glassware, large volumes of solutions, technical skills, and a long period of time. Using a similar principle for titration, we developed a novel µPAD that needs no calibration curve to accomplish acid-base titration.

This µPAD consists of reaction

reservoirs and detection reservoirs that contain varying amounts of a primary standard substance and a constant amount of a visible indicator.

A sample solution is dropped onto

the center of the µPAD, and is then allowed to spread to each reaction reservoir where it is neutralized by different amounts of the primary standard substance. The neutralized sample changes the color of the indicator in the detection reservoir only when the primary standard substance is insufficient to neutralize the sample solution.

The titration is

completed within one minute, including the time required for sample application and for determining the end point.

Therefore, this µPAD is more advantageous than classic

titrimetry in terms of speed, portability, and disposability, although the number of significant figures is less.

The design of this µPAD, and the volumes of the primary

standard, indicator, and sample solutions to be added were optimized to obtain reproducible results.

In addition, the stability of this µPAD was evaluated by storage

under various conditions of temperature and light exposure. We also used this µPAD for 6


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the on-site analysis of acidic water obtained from a natural hot spring.

EXPERIMENTAL SECTION Materials.

All reagents were of analytical grade and were used as received.

Deionized

water was prepared by means of an Elix water purification system (Millipore Co. Ltd., Molsheim, France).

Potassium hydrogen phthalate (KHPth), sodium hydroxide,

hydrochloric acid, nitric acid, sulfuric acid, 1 M sodium hydroxide solution (factor, 1.000), 1 M hydrochloric acid (factor, 1.001), phenolphthalein, orange I, and ethanol were purchased from Wako Pure Chemical Industries (Osaka, Japan).

Sodium carbonate,

acetic acid, and ammonia were obtained from Kanto Chemical (Tokyo, Japan). Bromocresol purple was purchased from Tokyo Kasei Kogyo (Tokyo, Japan).

Stock

solutions of 1 M KHPth and 1 M sodium carbonate were prepared by dissolving the appropriate amounts in water.

Stock solutions of 1(w/w)% phenolphthalein and

bromocresol purple were prepared by dissolving 1 g each of phenolphthalein and bromocresol purple in separate containers of 99 g of ethanol. µPAD design and fabrication.

We used Microsoft Office Power Point 2010 to design a

µPAD with a sample reservoir located at the center and ten reaction and detection reservoirs each arranged radially in a 30 × 30 mm square.

Details of the design of the

µPAD are provided in the Supporting Information (Figure S1). 7


According to the method


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reported by Carrilho and coworkers,7 the designed µPADs were printed on a sheet of filter paper (200 × 200 mm, Chromatography Paper 1CHR, WhatmanTM, GE Healthcare Lifesciences, United Kingdom) by a wax printer (ColorQube 8570N, Xerox, CT), followed by heating at 150 ºC for 2 min in a drying machine (ONW-300S, AS ONE Corporation, Osaka, Japan).

The back side of the printing surface was covered with clear packing tape

to prevent solution from leaking out underneath the µPAD, as reported by Mentele and

coworkers.13 The areas of the channels and reservoirs were measured using the public domain software Image J (National Institutes of Health).

Each µPAD was cut to a 30 ×

30 mm piece, and then was prepared by applying appropriate volumes of primary standard solutions and an indicator solution to the reaction reservoirs and detection reservoirs via a micropipette.

After drying, the µPAD was sandwiched by an acrylic plate holder that was

composed of two 30 × 30 mm plates.

The bottom plate had four stops that fix the top

plate, whereas the top plate had a hole (3 mmφ) to introduce a sample solution into the µPAD (Supporting Information, Figure S2).

An aliquot of a sample solution was gently

introduced into the µPAD through the hole in the top plate by dropping 30 µL of a sample solution via a micropipette.

Between titrations, the acrylic plate holder was flushed with

deionized water and then was wiped with disposable paper wiper. Analysis of acidic hot spring water.

An acidic hot spring water sample was obtained at

the source of the Tsukahara Onsen hot spring, Yufu, Oita, Japan. 8


Hot spring water was

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taken from a pool near a spa (40 ºC).

The hot spring water was passed through a

cation-exchange solid phase extraction (SPE) cartridge (HyperSep™ SCX Strong Cation Exchanger SPE Cartridge, 100 mg, Thermo Scientific, Thermo Fisher Scientific, MA) to remove metal ions and then was determined by the µ-PADs in the field.

To exchange H+

with Na+ before use, the cation-exchange SPE cartridge was pretreated with 1 mL of 1 M NaCl.

After the pretreatment, 0.25 mL of the hot spring water was added to the

cation-exchange SPE cartridge for flushing out the residual NaCl solution, and then 1.5 mL of the hot spring water was introduced to collect a sample for determination using the µ-PAD.

The cation-exchange SPE cartridge could be regenerated by flushing

successively with 1 mL of 5 M NaCl and 1 mL of 1 M NaCl and could be used at least five times. titration.

The hot spring water was also brought back to our laboratory for classic acid-base In the laboratory, the hot spring water was pretreated by the same procedure as

that carried out in the field analysis.

A 1-mL aliquot of the collected sample was titrated

with 0.01 M NaOH (factor, 1.000) using a 25-mL burette.

RESULTS AND DISCUSSION Applied volumes and concentrations of reagents.

A µPAD for acid-base titration

was prepared by applying primary standard solutions of acid or base in varying concentrations and a solution of an indicator, phenolphthalein, to the reaction and detection 9



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reservoirs.

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The volumes needed to fill the reaction and detection reservoirs were initially

estimated in order to optimize the applied volumes of the reagents.

In the preliminary

study, we confirmed that the volume needed for wetting the whole channel was 30 µL, according to a method reported by Dungchai and coworkers wherein different volumes of a colored solution were applied to the µPADs to fill the whole channel.28

Then, the

volumes used to fill the reaction and detection reservoirs were estimated by measuring the area of the whole channel, and those of the reaction and detection reservoirs.

The areas

were measured using Image J software (shown in Supporting Information, Table S1). The volumes of the reaction and detection reservoirs were obtained by multiplying the volume required to fill the whole channel by the area ratio of each reservoir in the whole channel.

The volumes needed to occupy the reaction and detection reservoirs were

calculated as 0.98±0.08 and 0.54±0.001 µL, respectively.

Therefore, 1 µL each of the

primary standard solutions was added to each reaction reservoir so as to be equivalent to the volume of the sample solution needed to fill the reaction reservoir, and 0.5 µL of a phenolphthalein solution was applied to the detection reservoirs to detect the excess amounts of acid or base that were not neutralized by the primary standard substance in the reaction reservoirs. It should be noted that the amount of phenolphthalein was an important factor in a clear visualization of a color change.

Therefore, concentrations of phenolphthalein that 10


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varied from 0.01 to 1% were added to the detection reservoirs without the addition of acid to the reaction reservoirs, and 0.1 and 0.01 M NaOH solutions were applied to the center of the µ-PAD in order to determine the optimal concentration for a clear visualization of color change, as shown in Figure 1.

Figure 1a shows the immediate color change with

different intensities in all the detection reservoirs that the phenolphthalein underwent when 0.10 M NaOH solution reached each of them. However, as the concentration decreased, the color gradually disappeared (Figure 1b). A concentration higher than 0.25% was needed for 0.1 M NaOH in order to determine a visible color change after the µ-PAD had dried.

Initially, the color could be seen in the detection reservoirs containing more than

0.25% of phenolphthalein for 0.01 M NaOH (Figure 1c), although it had almost disappeared in all the detection reservoirs after drying (Figure 1d).

Compared to Figure

1a, the color intensity in Figure 1c was obviously reduced, so that an increase in the concentration of phenolphthalein seemed better when using 0.01 M NaOH.

Consequently,

0.25% of phenolphthalein was employed for the titration of 0.1 to 1 M NaOH whereas the concentration of phenolphthalein was increased to 0.5% in the µPAD for measuring the samples containing from 0.01 to 0.1 M NaOH. With this system, we also could estimate the excess amounts of NaOH when the concentration of NaOH was higher than that of the primary standard added to the reaction reservoirs.

Assuming that the solution in the reaction reservoir was roughly 11



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homogeneous when neutralization progressed, the concentration of the NaOH solution penetrating into the detection reservoir was simply given by subtracting the concentration of the acid solution added to each reaction reservoir from that of the NaOH since the volume of the acid solution was equal to the sample volume that was needed to fill the reaction reservoir.

For example, when the µ-PAD contained 0.1 to 1 M of acid solution in

the reaction reservoirs, we can determine NaOH solutions with concentrations of 0.1−1 M. In the µ-PAD, the concentrations of hydroxide ion in the reaction reservoirs were more than 0.1 M after neutralization if the acid concentration in the reaction reservoir was lower than that of the NaOH solution; COH, detect = COH, sample − CH, react ≥ 0.1, where COH, detect, COH, sample,

and CH, react are the concentrations of OH− penetrating into the detection reservoir,

OH− in the sample solution, and the acid in the reaction reservoir, respectively.

Thus, the

concentration of the NaOH was sufficient to change the color of the phenolphthalein once it had penetrated the detection reservoir.

The situation was similar when using the µ-PAD

for 0.01 to 0.1 M NaOH; namely, the concentrations of NaOH were higher than 0.01 M when the concentrations of the acid were lower than that of NaOH. Acid-base titration.

To determine the concentrations of NaOH solutions, 1 µL of KHPth

solutions that varied from 0.1 to 1 M were applied to the ten reaction reservoirs in intervals of 0.1 M, whereas the detection reservoirs contained 0.5 µL of 0.25% phenolphthalein, which were the optimal conditions, as shown in Figure 1.

Thus, the µPADs permitted the

12


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determination of solutions with concentrations of 0.1-1 M NaOH. and 0.8 M NaOH solutions are shown in Figure 2.

The results for 0.4, 0.6,

The numbers on the µPAD indicate

the molar concentration of KHPth being added to the reaction reservoirs, as seen in Figure 2.

Each titration was completed within 1 min, which includes the time needed to apply

the sample solution (Supporting Iinformation, Video S1).

The detection reservoirs turned

to a red color when the concentrations of KHPth were lower than that of NaOH. Therefore, a clear red color was observed in detection reservoir numbers 0.3, 0.5, and 0.7 for 0.4, 0.6, and 0.8 M NaOH, respectively, as shown in Figures 2a, 2c, and 2e after the whole channel was occupied by the sample solution.

As the results show, the

concentrations of NaOH were determined directly with no calibration since each reaction reservoir contained a known amount of the primary standard substance, which neutralized the NaOH.

In practice, the concentrations of NaOH ranged from 0.31-0.40, 0.51-0.60,

and 0.71-0.80 M, as shown in Figures 2a, 2c, and 2e, respectively.

Reproducibility of the

µPAD was excellent since the same results were obtained for five independent measurements for each sample. The color is clearly shown in Figures 2a, 2c, and 2e where the µ-PADs were still wet, whereas the color change was unclear after drying, as shown in Figures 2b, 2d, and 2f. Furthermore, some detection reservoirs underwent an incorrect color change after drying for 60 min due to an unfavorable diffusion of excess hydroxide ion. 13


These results


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indicate that the end point should be found as soon as the sample solution occupies the whole channel. To determine lower concentrations of NaOH solutions, the standard solutions of KHPth were diluted 10-fold.

The solutions of KHPth with concentrations of 0.01 to 0.1

and 0.5% phenolphthalein were applied to the reaction and the detection reservoirs, and then 30 µL of 0.04, 0.06, and 0.08 M NaOH solutions were applied to the µPADs.

Clear

end points were found for dilute alkaline solutions with concentrations ranging from 0.01 to 0.1 M (Supporting Information, Figure S3). According to these results, the µPADs are, in fact, useful for the determination of NaOH at concentrations ranging from 0.01 M to 1 M since it is too difficult to measure these concentrations via conventional pH test paper (Supporting Information, Figure S4). We also fabricated the µPAD for measuring 0.005 to 0.05 M NaOH by adding 1% phenolphthalein to the detection reservoirs.

The results

showed that the µPAD was applicable to the determinations of 0.005 to 0.05 M NaOH at intervals of 0.005 M. Furthermore, by coupling two types of µPADs containing 0.1 to 1 M KHPth and 0.01 to 0.1 M KHPth, the concentrations of NaOH were determined more precisely at intervals of 0.02 M.

Initially, a sample was applied to the µPAD for 0.1 M to 1 M (high

concentration) to determine a rough concentration.

Then, the sample was mixed with an

equal volume of KHPth with the concentration determined by the first titration, followed 14


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by an application to the µPAD at 0.01 M to 0.1 M (low concentration).

Examples are

shown in Figure 3 where 0.25, 0.27, and 0.29 M NaOH were determined using the µ-PADs for both high and low concentrations.

The results shown in Figure 3a were obtained by

the µ-PAD for high concentrations and show that the sample contained NaOH in concentrations ranging from 0.21 to 0.30.

Then, we added an equal volume of 0.20 M

KHPth and applied the mixture to the µ-PAD for low concentrations.

The results shown

in Figure 3b, which were obtained by the µPAD for low concentrations, suggest that the concentration of the mixture was from 0.021 to 0.030 M. Consequently, it is apparent that the concentration of NaOH in the sample ranged from 0.242 to 0.260, since the sample solution was diluted two-fold before being applied to the µ-PAD for low concentrations. For 0.27 M and 0.29 M NaOH, the µPAD for high concentrations (0.1-1 M) showed the same results (Figure 3c and 3e) as that of 0.25 M with good reproducibility using triplicate measurements, as expected, whereas the mixtures with 0.2 M KHPth were determined to be from 0.031 to 0.040 M for 0.27 M NaOH (Figure 3d) and from 0.041 to 0.050 M for 0.29 M NaOH (Figure 3f).

Therefore, we should be able to discriminate these

concentrations by adding a certain amount of KHPth to the sample, followed by a measurement using the µPAD for low concentrations (0.01-0.1 M) The same titration method is applicable to the determination of HCl by using Na2CO3 as the primary standard substance instead of KHPth.

In this case, when the reaction

15



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reservoirs contain Na2CO3 at concentrations higher than that of the sample solution (acid), a red color is observed in the corresponding detection reservoir.

The results for HCl

solutions for concentrations of 0.4, 0.6, and 0.8 M are shown in Figure 4.

The

performances of µPADs for HCl were similar to those for NaOH, so both acid and base were determined with only the use of an appropriate primary standard substance. We also determined nitric acid, sulfuric acid, acetic acid, and ammonia since the µ-PAD is expected to be applicable to the determination of other acids and bases.

Strong

acids (nitric acid and sulfuric acid) and a weak acid (acetic acid) were successfully determined as well as HCl with good reproducibility.

The µ-PAD also worked well with

a clear end point in the determination of ammonia solutions with the concentrations of 0.1 to 1 M, although the intensity of the red color was lower than that obtained by NaOH due to the weak basic property of ammonia.

However, the end point for 0.01 to 0.1 M

ammonia was undetectable as shown in Figure 5a where 0.07 M of an ammonia solution was determined using KHPth and phenolphthalein as the acid and indicator.

As Figure 5a

shows, we could not find the correct end point due to the weak intensity of the color, although the detection reservoirs of 0.01 to 0.05 seemed to be slightly red.

However, the

detection reservoir for 0.06 M must turn a definite red color since the concentration of ammonia is 0.07 M.

In addition, the slight red color had completely disappeared within a

few minutes after completing the titration.

Ammonia is a weak base with a pKb of 4.76 16


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so that the pH of 0.01 M ammonia solution was estimated to be 10.6.

A pH of 10.6 was

much lower than that of 0.01 M NaOH (pH=12), resulting in a difficulty of visual detection. To detect the end point for diluted ammonia solutions, a suitable indicator should be selected instead of phenolphthalein.

Methyl orange may be a candidate for detecting an

excess amount of ammonia since its indicator range is acidic (3.1~4.4).

However, it was

difficult to find the color change from orange to yellow on the µ-PAD with the naked eyes. Consequently, to detect ammonia at a low concentration, we employed bromocresol purple as the indicator, because it has an indicator range (5.2~6.8) that is lower than that of phenolphthalein (8.3~10) and the color change from yellow to purple is clearer than that from orange to yellow. Figure 5b shows the results of 0.07 M ammonia using bromocresol purple as the indicator.

A clear end point at 0.07 M was found in Figure 5b where the end point

became more visible than that in Figure 5a.

These results indicate that several weak acids

and weak bases can be determined by selecting an appropriate indicator to detect the end point. Stability of µPADs.

The developed µPADs can definitely be employed for point-of-use

and rapid measurements of acid and base concentrations for field analysis.

However, the

µPADs must have a long lifetime for conventional use in the field where no 17



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instrumentation is available.

Page 18 of 32

Therefore, we examined the stability of the µPADs under

conditions of different temperature and light exposures. When the µPADs were stored at room temperature under light, they only worked for five days.

After five days, the phenolphthalein showed an unclear color change.

Conversely, when we kept the µPADs at room temperature and a low temperature of 4 ºC in the dark, they were effective for more than a month.

Therefore, the µPADs should be

stored in the dark before use in order to obtain reproducible results.

The instability can

obviously be attributed to the susceptibility of phenolphthalein to light, as indicated in the safety data sheet. On-site analysis. To demonstrate the utility of the µ-PAD in on-site analysis, acidic hot spring water was determined at the site of the Tsukahara Onsen hot spring.

The hot

spring water contained a high concentration of iron ion which formed precipitation of hydroxide by consuming hydroxide ion, resulting in a positive error.

So, in the analysis,

the iron ion was removed using a cation-exchange SPE cartridge before the titration. Figure 6a shows the tools used for the titration by the µ-PAD in the field. noted that no glassware was needed for the titration by the µ-PAD.

It should be

The results obtained

in the field by the µPAD for 0.005 to 0.05 M are shown in Figure 6b.

The acid

concentration of the hot spring water was determined to be 0.020 M (practically ranging from 0.020 to 0.024 M) with good reproducibility in triplicate measurements by the µPAD. 18


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The hot spring water was also determined at the laboratory by classic titration, and the results showed a concentration of 0.0222±0.0005 M (n=5) which was consistent with those of the on-site analysis.

Therefore, the µ-PAD permitted acid-base titration in the

field without using glassware.

CONCLUSIONS A simple and rapid titration method was developed using µ-PADs fabricated by a wax printer.

The newly designed µ-PADs were successfully applied to the titrations of both

acids and bases using KHPth and Na2CO3 as the primary standard substances, respectively. Titration was completed within one minute by dropping 30 µL of a sample solution via a micropipette.

The µ-PAD has advantages in portability and disposability since it is small,

light, and inexpensive.

It should be emphasized that the method needs no calibration

curve since the concentration of the sample is determined directly by the known amounts of a primary standard substance that has been added to the reaction reservoirs.

The

µPAD permitted acid-base titration in the field as demonstrated in the analysis of an acidic hot spring water sample.

A similar strategy could be applicable to the other titration

methods including chelate titration, redox titration, and precipitation titration by using an appropriate primary standard substance and an indicator.

Consequently, the proposed

µ-PAD is a viable alternative to classic titration methods in the analysis of several 19



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chemical species in the field at sites where glassware and large volumes of solutions are difficult to obtain.

Corresponding Author *E-mail: [email protected]. Tel: +81-86-251-7847. Fax: +81-86-251-7847. Notes The authors declare no competing financial interest.

ACKNOWLEDGMENTS This research was supported by The Yakumo Foundation for Environmental Science and Grants-in-Aid for Scientific Research, Scientific Research (B) (No. 26288067).

We

would like to thank Ms. M. Sakurai, Mr. T. Shinohara, Mr. T. Abe, and Mr. H. Takahata of Beppu Hakudo Kogyo Corp. for their kind help in the sampling of the hot spring water.

SUPPORTING INFORMATION AVAILABLE Estimated volumes of solution needed to fill the µ-PAD, the design of the µPAD, a photo of an acrylic plate holder, titration of NaOH solutions with concentrations of 0.04, 0.06, and 0.08 M, color change of conventional pH test paper, and a video of titration. information is available free of charge via the Internet at http://pubs.acs.org/.

20


This

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REFERENCES (1) Lagally, E. T.; Medintz, I.; Mathies R. A. Anal. Chem. 2001, 73, 565–570. (2) Jacobson, S. C.; Hergenroder, R); Koutny, L. B.; Ramsey, J. M. Anal. Chem. 1994, 66, 1114–1118. (3) Lee, J. H.; Song, Y.-A.; Tannenbaum, S. R.; Han, J. Anal. Chem. 2008, 80, 3198–3204. (4) Qin, D.; Xia, Y.; Rogers, J. A.; Jackman, R. J.; Zhao, X.; Whitesides, G. M. Top. Curr. Chem. 1998, 194, 1−20. (5) Martinez, A. W.; Phillips, S. T.; Butte, M. J.; Whitesides, G. M. Angew. Chem., Int. Ed. 2007, 46, 1318−1320. (6) Lu, Y.; Shi, W.; Jiang, L.; Qin, J.; Lin, B. Electrophoresis 2009, 30, 1497–1500. (7) Carrilho, E.; Martinez, A. W.; Whitesides, G. M. Anal. Chem. 2009, 81, 7091−7095. (8) Maejima, K.; Tomikawa, S.; Suzuki, K.; Citterio, D. RSC Adv. 2013, 3, 9258−9263. (9) Dungchai, W.; Chailapakul, O.; Henry, C. S. Analyst 2011, 136, 77−82. (10) Cassano, C. L.; Fan, Z. H. Microfluid. Nanofluid. 2013, 15, 173−181. (11) Liu,W.; Cassano, C. L.; Xu, X,; Fan, Z. H. Anal. Chem. 2013, 85, 10270−10276. (12) Zhang, Y.; Zhou C.; Nie, J.; Le, S.; Qin, Q.; Liu, F.; Li, Y.; Li, J. Anal. Chem. 2014, 86, 2005−2012. (13) Mentele, M. M.; Cunningham, J.; Koehler, K.; Volckens, J.; Henry, C. S. Anal. Chem. 2012, 84, 4474−4480. 21



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(14) Rattanarat, P.; Dungchai, W.; Cate, D. M.; Siangproh, W.; Volckense, J., Orawon Chailapakul, O.; Henry, C. S. Anal. Chim. Acta 2013, 800 50– 55. (15) Cate, D. M.; Nanthasurasak P.; Riwkulkajorn, P.; L’Orange, C.; Henry, C. S.; Volckens, J. Ann. Occup. Hyg. 2014, Vol. 58, 413–423. (16) Yamada, K.; Takaki, S.; Komuro N.; Suzuki, K.; Citterio, D. Analyst 2014, 139, 1637–1643. (17) Li, X.; Tian, J.; Shen, W. Anal. Bioanal. Chem. 2010, 396, 495–501. (18) Wang, Y; Wang, S.; Ge, S.; Wang, S.; Yan, M.; Zang, D.; Yu, J. Monatsh. Chem. 2014, 145, 129–135 (19) Jokerst, J. C.; Adkins, J. A.; Bisha, B.; Mentele, M. M.; Goodridge, L. D.; Henry, C. S. Anal. Chem. 2012, 84, 2900−2907. (20) Cheng, C.-M.; Martinez, A. W.; Gong, J.; Mace, C. R.; Phillips, S. T.; Carrilho, E.; Mirica, K. A.; Whitesides, G. M. Angew. Chem., Int. Ed. 2010, 122, 4881−4884. (21) Rattanarat, P.; Dungchai, W.; Siangproh, W.; Chailapakul, O.; Henry, C. S. Anal. Chim. Acta 2012, 744, 1−7. (22) Lankelma, J.; Nie, Z.; Carrilho, E.; Whitesides, G. M. Anal. Chem. 2012, 84, 4147−4152. (23) Santhiago, M.; Wydallis, J. B.; Kubota, L. T.; Henry, C. S. Anal. Chem. 2013, 85, 5233−5239. (24) Santhiago, M.; Kubota, L. T. Sens. Actuators, B 2013, 177, 224– 230. 22


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(25) Cate, D. M.; Dungchai, W.; Cunningham, J. C.; Volckens J.; Henry C. S. Lab Chip 2013, 13, 2397−2404. (26) Lewis, G. G.; Robbins, J. S.; Phillips, S. T. Anal. Chem. 2013, 85, 10432–10439. (27) Lewis, G. G.; Robbins, J. S.; Phillips, S. T. Chem. Commun. 2014, 50, 5352–5354. (28) Dungchaia, W.; Chailapakul, O; Henry, C. S. Anal. Chim. Acta 2010, 674, 227–233.

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FIGURE LEGENDS Figure 1.

Optimization of the concentration of phenolphthalein added to the detection

reservoirs. The concentration of phenolphthalein; Reservoir number, 1=0.01% , 2=0.025%, 3=0.050%, 4=0.075%, 5=0.10%, 6=0.25%, 7=0.50%, 8=0.75%, 9=1%, 10=0% (blank); volume added, 0.5 µL.

(a) Immediate image after the addition of 0.1 M NaOH, (b) image of (a) after

drying, (c) immediate image after the addition of 0.01 M NaOH, (d) image of (c) after drying.

Figure 2.

Titration of NaOH solutions with different concentrations.

(a) 0.4 M NaOH

(immediately after addition), (b) 0.4 M NaOH (after drying), (c) 0.6 M NaOH (immediately after addition), (d) 0.6 M NaOH (after drying), (e) 0.8 M NaOH (immediately after addition), (f) 0.8 M NaOH (after drying).

The numbers of the

reservoirs indicate the concentrations of KHPth solutions added to the reaction reservoirs. Detection reservoirs contained 0.5 µL of 0.25% phenolphthalein.

Figure 3.

Titration of 0.25, 0.27, and 0.29 M NaOH solutions.

The numbers of the

reservoirs indicate the concentrations of KHPth solutions added to the reaction reservoirs. (a) Detection reservoirs, 0.5 µL of 0.25% phenolphthalein; sample, 0.25 M NaOH. 24


(b)

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Detection reservoirs, 0.5 µL of 0.5% phenolphthalein; sample, a mixture of equal volumes of 0.25 M NaOH and 0.20 M KHPth that should contain 0.025 M of hydroxide ion.

(c)

Detection reservoirs, 0.5 µL of 0.25% phenolphthalein; sample, 0.27 M NaOH.

(d)


(e)

Detection reservoirs, 0.5 µL of 0.25% phenolphthalein; sample, 0.29 M NaOH.

(f)


Figure 4.

Titration of HCl solutions with different concentrations.

The numbers of the reservoirs indicate the concentrations of Na2CO3 solutions added to the reaction reservoirs.

Detection reservoirs contained 0.5 µL of 0.5% phenolphthalein.

(a) 0. 4 M HCl, (b) 0.6 M HCl, (c) 0.8 M HCl.

Figure 5.

Titration of 0.07 M ammonia solution.

Indicator, (a) 0.5% phenolphthalein, (b) 0.5% bromocresol purple. reservoirs contained 1 µL of 0.01 to 0.1 M KHPth solutions. contained 0.5 µL of the indicator.

25


The reaction

Detection reservoirs


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Figure 6.

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Titration of water samples from the source of a hot spring in the field.

(a) Tools for on-site analysis.

1=disposable syringe for applying pressure to a

cation-exchange SPE cartridge, 2=cation-exchange SPE cartridge, 3=sample tube, 4=µ-PAD, 5=acrylic plate holder.

(b) Titration by the µ-PAD.

contained 1 µL of 0.005 to 0.050 M Na2CO3 solutions. 0.5 µL of 1% phenolphthalein.

The reaction reservoirs

Detection reservoirs contained

The insert shows the sampling from the source of a hot

spring.

26


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Figure 1. Optimization of the concentration of phenolphthalein added to the detection reservoirs. The concentration of phenolphthalein; Reservoir number, 1=0.01% , 2=0.025%, 3=0.050%, 4=0.075%, 5=0.10%, 6=0.25%, 7=0.50%, 8=0.75%, 9=1%, 10=0% (blank); volume added, 0.5 µL. (a) Immediate image after the addition of 0.10 M NaOH, (b) image of (a) after drying, (c) immediate image after the addition of 0.01 M NaOH, (d) image of (c) after drying. 926x1190mm (96 x 96 DPI)



1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Figure 2. Titration of NaOH solutions with different concentrations. (a) 0.4 M NaOH (immediately after addition), (b) 0.4 M NaOH (after drying), (c) 0.6 M NaOH (immediately after addition), (d) 0.6 M NaOH (after drying), (e) 0.8 M NaOH (immediately after addition), (f) 0.8 M NaOH (after drying). The numbers of the reservoirs indicate the concentrations of KHPth solutions added to the reaction reservoirs. Detection reservoirs contained 0.5 µL of 0.25% phenolphthalein. 926x1190mm (96 x 96 DPI)


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Figure 3. Titration of 0.25, 0.27, and 0.29 M NaOH solutions. The numbers of the reservoirs indicate the concentrations of KHPth solutions added to the reaction reservoirs. (a) Detection reservoirs, 0.5 µL of 0.25% phenolphthalein; sample, 0.25 M NaOH. (b) Detection reservoirs, 0.5 µL of 0.5% phenolphthalein; sample, a mixture of equal volumes of 0.25 M NaOH and 0.20 M KHPth that should contain 0.025 M of hydroxide ion. (c) Detection reservoirs, 0.5 µL of 0.25% phenolphthalein; sample, 0.27 M NaOH. (d) Detection reservoirs, 0.5 µL of 0.5% phenolphthalein; sample, a mixture of equal volumes of 0.27 M NaOH and 0.20 M KHPth that should contain 0.035 M of hydroxide ion. (e) Detection reservoirs, 0.5 µL of 0.25% phenolphthalein; sample, 0.29 M NaOH. (f) Detection reservoirs, 0.5 µL of 0.5% phenolphthalein; sample, a mixture of equal volumes of 0.29 M NaOH and 0.20 M KHPth that should contain 0.045 M of hydroxide ion. 381x457mm (96 x 96 DPI)



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Figure 4. Titration of HCl solutions with different concentrations. The numbers of the reservoirs indicate the concentrations of Na2CO3 solutions added to the reaction reservoirs. Detection reservoirs contained 0.5 µL of 0.5% phenolphthalein. (a) 0. 4 M HCl, (b) 0.6 M HCl, (c) 0.8 M HCl. 381x457mm (96 x 96 DPI)


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Figure 5. Titration of 0.07 M ammonia solution. Indicator, (a) 0.5% phenolphthalein, (b) 0.5% bromocresol purple. The reaction reservoirs contained 1 µL of 0.01 to 0.1 M KHPth solutions. Detection reservoirs contained 0.5 µL of the indicator. 381x457mm (96 x 96 DPI)



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Figure 6. Titration of water samples from the source of a hot spring in the field. (a) Tools for on-site analysis. 1=disposable syringe for applying pressure to a cation-exchange SPE cartridge, 2=cation-exchange SPE cartridge, 3=sample tube, 4=µ-PAD, 5=acrylic plate holder. (b) Titration by the µ-PAD. The reaction reservoirs contained 1 µL of 0.005 to 0.050 M Na2CO3 solutions. Detection reservoirs contained 0.5 µL of 1% phenolphthalein. The insert shows the sampling from the source of a hot spring. 381x457mm (96 x 96 DPI)


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