An electricity- and instrument-free infectious disease sensor based on a 3D origami paper-based analytical device

Chung-An Chen a, Hao Yuan a, Chiao-Wen Chen a, Yuh-Shiuan Chien a, Wang-Huei Sheng b and Chien-Fu Chen *a
aInstitute of Applied Mechanics, National Taiwan University, Taipei 106, Taiwan. E-mail:
bDivision of Infectious Diseases, Department of Internal Medicine, National Taiwan University Hospital, Taipei 100, Taiwan

Received 1st February 2021 , Accepted 29th March 2021

First published on 30th March 2021

Infectious diseases cause millions of deaths annually in the developing world. Recently, microfluidic paper-based analytical devices (μPADs) have been developed to diagnose such diseases, as these tests are low cost, biocompatible, and simple to fabricate. However, current μPADs are difficult to use in resource-limited areas due to their reliance on external instrumentation to measure and analyze the test results. In this work, we propose an electricity and external instrumentation-free μPAD sensor based on the colorimetric enzyme-linked immunosorbent assay (ELISA) for the diagnosis of infectious disease (3D-tPADs). Designed based on the principle of origami, the proposed μPAD enables the sequential steps of the colorimetric ELISA test to be completed in just ∼10 min. In addition, in order to obtain an accurate ELISA result without using any instrument, we have integrated an electricity-free “timer” within the μPAD that can be controlled by the buffer viscosity and fluid path volume to indicate the appropriate times for washing and color development steps, which can avoid false positive or false negative results caused by an extended or shortened amount of washing and development times. Due to the low background noise and high positive signal intensity of the μPAD, positive and negative detection results can be distinguished by just the naked eye. Furthermore, the ELISA result can be semi-quantified by comparing the results shown on the μPAD with a color chart diagram with a detection limit of HIV type 1(HIV-1) p24 antigen as low as 0.03 ng mL−1. These results demonstrate the proposed sensor can perform infectious disease diagnosis without external instrumentation or electricity, extending the application of the μPAD test for on-site detection and use in resource-limited settings.

1. Introduction

Infectious diseases caused by pathogenic microorganisms (e.g., bacteria, viruses, parasites, and fungi) are currently the second leading cause of death worldwide, particularly in low- and middle-income nations.1–5 For example, acquired immunodeficiency syndrome (AIDS) is caused by the human immunodeficiency virus (HIV) and is the leading cause of death in South Africa.1,2 Accurate and rapid diagnosis is essential to manage the spread of this infectious disease and can help with better clinical decision-making as well as disease control and prevention.3 However, current testing methods, such as culture- and nucleic-acid-based techniques,4,5 are often unavailable in the developing world due to resource limitations.

To address this issue, lateral flow assays (LFAs), which are a type of paper-based device that can perform rapid immunoassays via capillary action of the reagents, have been proposed for pathogen diagnosis in developing countries as they are low-cost, relatively simple to manufacture, do not require power, and can be made widely available.6,7 Most LFAs allow analytes to be transported through various detection media and result in a simple to use colorimetric or fluorescent readout.8–11 However, LFAs lack the functionality to conduct multiple reaction steps that could provide an all-in-one sample-to-answer solution for disease diagnosis.12 Additionally, LFAs generally possess limited sensitivity and can only provide qualitative detection results.12

To enhance the sensitivity and enable multi-step and quantitative detection, three dimensional (3D) microfluidic paper-based analytical devices (μPADs) have been developed, enabling sequential steps of assays to be easily performed by stacking and folding multiple fluidic layers from a single sheet of paper.13–16 This design provides 3D μPADs with superior advantages over LFAs in terms of sensitivity, multiplexity, sample consumption, and quantitative analysis, with strong potential for point-of-care disease diagnosis.7,17–23 In these devices, pathogenic microorganisms are detected through various means including electrochemical or fluorescent signaling.24–27 Although such 3D μPADs are simple, integrated, and highly miniaturized, external instruments (e.g., smart phones or excitation light sources) are still required to complete the assays. Additionally, the operation of these devices depends on electricity, making them impractical to use in the field or resource-limited areas of the world.

To address these obstacles, we demonstrate an electricity- and external instrument-free sensor for infectious disease diagnosis based on a 3D μPAD. By performing a colorimetric enzyme-linked immunosorbent assay (ELISA) on the μPAD, the presence of the target analyte can be converted to a dark blue color that is visible to the naked eye, preventing the need for an excitation light source or smart phone. The 3D μPAD adopts the principle of origami (paper folding), enabling the multi-step ELISA test to be conducted by simply folding and sliding the paper-based components of the μPAD. As a result, the assay can be easily performed, even by non-experts. Additionally, we have integrated an electricity-free “timer” into the 3D-tPADs, which is functionalized by delivering colored buffer to certain wells on the 3D-tPADs at the appointed times. By adjusting the viscosity of buffer and the volume of fluid path, the buffer can be delivered at the appropriate timing for each ELISA step. The integration of this timer improves the user experience by allowing untrained personnel to perform the assay, making the proposed 3D-tPADs to conform to the ASSURED (affordable, sensitive, specific, user-friendly, rapid and robust, equipment-free and deliverable to end-users) criteria set by the World Health Organization (WHO) for point-of-care tests.

To verify the feasibility of the sensor for infectious disease diagnosis, we detect blood plasma containing HIV type 1 (HIV-1) p24 antigen, which is recognized as a virological biomarker in early HIV detection.28 The presence of HIV-1 p24 is indicated by a color change on the 3D-tPADs, which features a detection limit as low as 0.03 ng mL−1 due to the low background noise and high positive signal intensity. Furthermore, the HIV-1 p24 concentration can be semi-quantified by comparing the resulting test color on the 3D-tPADs with a pre-prepared color chart diagram, which shows the detection results based on different antigen concentrations. Throughout the whole detection process, no electricity nor external equipment is required, enabling the sensor to be applied for on-site infectious disease diagnosis and extending its usage to resource-limited areas and the developing world.

2. Experimental

2.1. Chemicals and reagents

HIV-1 p24/capsid protein p24 ELISA pair set (mouse anti-HIV-1 p24, recombinant HIV-1 p24, mouse anti-HIV-1 p24-HRP) was purchased from Sino Biological (Sino Biological, Beijing, China). Whatman® cellulose chromatography paper (grade 1 Chr and 3MM Chr sheets), phosphate buffered saline (PBS, pH 7.4), phosphate buffered saline containing TWEEN® 20 (PBS-T, pH 7.4), bovine serum albumin (BSA), 3,3′,5,5′-tetramethylbenzidine (T-MB) solution, sodium carboxymethyl cellulose (CMC), N-(3-di-methylaminopropyl)-N′-ethylcarbodiimide hydrochloride (ED-C), N-hydroxysuccinimide (NHS), and sodium acetate buffer solution (NaOAc buffer) were obtained from Sigma-Aldrich (Sigma-Aldrich, St. Louis). Sucrose and calcium chloride dehydrate were obtained from J. T. Baker® (Phillipsburg, NJ). Pullulan was purchased from Tokyo Chemical Industry. The green food dye was purchased from Youqi Co., Ltd. (Wugu, Taiwan). The ultrapure water was obtained from a Sartorius Arium® mini plus lab water system (Göttingen, Germany). The human blood samples used in this work were collected from a healthy volunteer using a vacutainer and approved by the institutional review board of Chang Gung Memorial Hospital, Taiwan.

2.2. Instruments

The HIV-1 p24 detection results on the chromatography paper were imaged using a smartphone (Apple iPhone, Cupertino, CA), and then analyzed with ImageJ software (National Institutes of Health, Bethesda, MD) to determine the red and blue values. A micro refrigerated centrifuge was purchased from KUBOTA (Kubota 3520, Tokyo, Japan) and used to separate the plasma from the blood samples. Solid wax hydrophobic barriers were deposited on the cellulose substrate using a wax printer (Xerox 8570, Fuji, Japan). The holder was fabricated using a 3D printer (Flashforge, Hong Kong, China).

2.3. Fabrication of the 3D-tPADs

The proposed 3D-tPADs was fabricated by folding a patterned sheet of cellulose chromatography paper based on the principle of origami (Fig. S1). The 3D-tPADs consisted of a strip, a detection pad, an indication pad, and 3 absorbent pads, as shown in Fig. S2. Each strip or pad of the 3D-tPADs consisted of hydrophilic channels, wells, or holes surrounded by hydrophobic barriers, which were fabricated by a previously developed wax printing method 2. Specifically, the solid wax was first patterned on the cellulose paper using the wax printer, and then heated at 120 °C for 1 min to help the wax penetrate between the cellulose fibers and form the hydrophobic barriers throughput the paper thickness on the 3D-tPADs. The holes on the strip were fabricated with a paper punch.

2.4. Surface modification of the 3D-tPADs

The detection wells on the detection pad of the 3D-tPADs (Fig. S2) were modified with capture antibodies (anti-HIV-1 p24), which were used to capture the HIV-1 p24 antigen in the sample. Specifically, 100 μL of 25 mM CaCl2 solution (pH 6) containing 0.5% (w/w%) CMC was first added onto the detection wells, and then rested at room temperature for 10 min, followed by drying at 40 °C for 5 min. Afterward, 100 μL of NaOAc buffer containing EDC (0.1 M) and NHS (0.4 M) was added to the CMC-treated detection wells and rested for 30 min, followed by drying at 40 °C for 5 min. 3 μL of the capture antibody was then added to the detection wells. The resulting 3D-tPADs was dried at room temperature, then folded (Fig. S1) and placed in a sealed bag and stored in a 4 °C refrigerator.

2.5. Reagent pre-drying on the 3D-tPADs

To prepare the ELISA reagents in the 3D-tPADs, we first added 3 μL of detection antibody (anti-HIV-1 p24-HRP) to the conjugate wells on the strip (Fig. S2) (without prior surface modification), which were then allowed to dry. This pre-dried detection antibody is used for binding with the capture antibody-absorbed HIV-1 p24 to form the antigen–antibody complex. Then we added 3 μL of green food dye to the dye wells (Fig. S2) on the detection pad, which were also allowed to dry. As a result, when fluid flows through the dye well, the dried green food dye can dissolve and color the fluid green. Finally, the hydrophilic channels and wells on the absorbent pads were treated with aqueous sucrose and pullulan solutions and dried to increase the viscosity of the sample fluid flowing though these channels and wells. The mass of the sucrose and pullulan in the wells were adjusted by tuning the deposition area on the absorption pads and the concentrations of sucrose and pullulan, as shown in Tables S1 and S2 in the ESI.

2.6. Optimization of the color development time

To optimize the color development time, we conducted an HIV-1 p24 detection test on the proposed 3D-tPADs. Prior to HIV-1 p24 detection, the folded 3D-tPADs was first secured in place using a 3D-printed sample holder (Fig. S3). To perform the HIV-1 p24 ELISA detection test using the 3D-tPADs, 3 μL of PBS containing 0, 0.01, 0.03, 0.1, 0.3, 1, 3, 10, or 30 ng mL−1 HIV-1 p24 was dropped through the holes located on the strip onto the T well, and 3 μL PBS containing no HIV-1 p24 was dropped onto the C well (Fig. S4(a)). After 1 min, we slid the strip down until the conjugate wells overlapped with the detection wells and added 100 μL of PBS-T with 5 wt% BSA (washing buffer) onto each conjugate well to rinse the dried detection antibody onto the T and C wells (Fig. S4(b)). After the washing buffer was completely absorbed by the underlying absorbent pads (around 4 min), we slid the strip down again until the holes on tab 3 overlapped with the T and C wells, to which we added 3 μL of the TMB solution (Fig. S4(c)). Then we imaged the resulting well at 1, 3, 5, 7 and 9 min, and analyzed the blue and red values of the T and C wells using ImageJ (Fig. S4(d)).

2.7. HIV-1 p24 detection process on the 3D-tPADs with the electricity-free timer

The HIV-1 p24 detection process followed the same procedures described in the preceding section, only utilizing the developed “timer”. The “timer” refers to the two wells on the indicator pad (labeled with “1” and “2”, as shown in Fig. S2). The timer is started when the washing buffer is added onto the T and C wells (Fig. S5), in which the buffer soaks into the underlying absorbent pads and is then delivered to the dye wells, where the dried green dye will be dissolved into the buffer. Ultimately, the green washing buffer will be delivered to the indication wells (Fig. S5) on the indicator pad, coloring these wells with green. By controlling the area and thickness of the absorbent pads and adjusting the viscosity of the washing buffer, the wells on the indicator pad can be colored at the 4th and 9th minutes and used as a timer to perform the multistep ELISA test on the proposed 3D-tPADs. Specifically, after triggering the timer by adding the washing buffer to the T and C wells, well 1 on the indicator pad will be colored green at the 4th minute. Therefore, at this time we added 3 μL of TMB solution to the T and C wells through the holes on tab 1. Then at the 9th minute, well 2 will turn green, which indicates the appropriate time at which the T and C wells should be imaged with a smartphone to analyze their blue and red values.

Following the same procedures, we detected blood plasma containing 0, 0.01, 0.03, 0.1, 0.3, 1, 3, 10, and 30 ng mL−1 HIV-1 p24 to investigate the feasibility of the proposed 3D-tPADs for clinical sample detection.

3. Results and discussion

3.1. The operation process and working principles of the proposed sensor

For the proposed disease sensor, we developed a 3D-tPADs that can perform the ELISA test, as shown in Fig. 1(a). Specifically, we designed the 3D-tPADs based on the principles of origami to minimize the manual operation steps and the device size. The 3D-tPADs (Fig. 1(b) and S2) is comprised of a detection pad featuring test (T) and control (C) detection wells, which are surface modified with capture antibodies used to absorb the antigens when the sample solution is added onto these wells (Fig. 1(b)). The 3D-tPADs also includes a strip of paper featuring two conjugate wells with dried detection antibodies (Fig. 1(b)), which are washed down to the detection wells after adding the washing buffer to the conjugate wells and bind with the antigens absorbed by the capture antibody on the detection wells. To incorporate an automated timer function in the device, we also create a fluidic path that delivers the excess washing buffer from the absorbent pads to the dye wells, which are pre-treated with dried green food dye, thus coloring the solution and turning the indication wells green. This color signals the time at which different steps in the assay should be performed (Fig. 1(b)). By pre-treating the paper substrate with dried sucrose and pullulan, we are able to control the viscosity, and therefore the flow rate, of the washing buffer to ensure it transfers the dye to the indication wells at the appropriate times for conducting different steps of the test.
image file: d1lc00079a-f1.tif
Fig. 1 (a) Images of the components of the proposed infectious disease sensor, including a 3D-tPADs, 3D-printed holder, and color chart diagram. (b) The structure of the proposed 3D-tPADs. The major components include (b1) conjugate wells pre-dried with detection antibodies, (b2) detection wells modified with capture antibodies, and (b3) absorbent pads with hydrophilic channels and wells. The channels and wells are dried with sucrose and pullulan, which dissolve into the washing buffer to increase its viscosity and lower its flow rate. (c) The operation procedure of the ELISA test on the proposed 3D-tPADs. (c1) First the test sample is added to the T well and the control sample (PBS) to the C well. In this step, the antigen contained in the test sample will be absorbed by the capture antibody in the T well, while no antigen will be captured by the C well (c2). (c3) Next, we slide the paper strip to expose the conjugate wells and add washing buffer. In this step, the HRP-conjugated antibody will be rinsed to the detection wells and form the antibody–antigen complex on the T well, while no complex will be formed on the C well (c4). Afterward, the redundant washing buffer will be soaked into the absorbent pads underlying the T and C wells to activate the timer, and then delivered to the dye wells, dissolving the pre-dried green food dye and ultimately delivering this color to the indication wells. (c5) After indication well 1 displays green color (i.e., serving as the timer function in the device), we slide the strip further to expose the C and T wells and add TMB solution. In the T well, the TMB solution will react with the HRP conjugated on the antibody and develop a dark blue color, while the C well displays a light blue color (c6). (c7) When indication well 2 displays green color, this indicates the appropriate time to compare the results shown on the T well with the prepared color scale diagram.

Based on the origami design of the proposed 3D-tPADs and the developed timer, the operation of the detector can be simplified into 4 steps:

(1) Sampling: first we add the test sample to the T well and control sample to the C well through the holes on tab 1 (Fig. 1(c1)), which have been pre-modified with capture antibodies (Fig. 1(b)). After adding the samples, the antigens contained in the test sample will bind with the capture antibodies and form the antigen-capture antibody complex on the T well. Meanwhile, for the C well, no antigen–antibody complex will form since no antigens are contained in the control sample (Fig. 1(c2)).

(2) Washing: next, we pull down the paper strip (Fig. S4) until the T and C wells overlap with the conjugate wells (on the orange tab), to which we add 100 μL of washing buffer (Fig. 1(c3)). The HRP-conjugated detection antibodies pre-dried on the conjugate wells (Fig. 1(b)) will be rinsed onto the T and C wells after adding the washing buffer and react with the antigen–antibody complex formed in step (1) on the T well (Fig. 1(c3 and c4)).

In addition to rinsing the antibodies, the addition of the washing buffer also activates the timer. Specifically, after being added onto the T and C wells, the excessive washing buffer soaks into the underlying absorbent pads (Fig. 1(b3)), where it dissolves the pre-dried sucrose and pullulan, and then being colored with green after melting the pre-dried green food dye on the dye wells, and ultimately delivering the green color to the indication wells to indicate the timing for performing the immunoassay steps as shown in Fig. S5.

(3) Color development: when indication well 1 shows green color (the time of which is determined after we optimized the reaction and fluidic flow rate), we slide down the strip, and add TMB solution to the T and C wells through the holes on tab 3 (Fig. 1(c5)). As a result, when the biomarker is present, the T well will display a dark blue color due to the reaction between the TMB solution and the HRP tagged on the detection antibody; while the C well will display a light blue color due to the reaction between TMB solution and HRP conjugated on the residual detection antibody that physically adsorbed on the C well, as shown in Fig. 1(c6).

(4) Comparing: when indication well 2 displays green color (again, the time of which was determined from optimization; Fig. 1(c4)), this indicates the optimal TMB and HRP reaction time. At this point in the test, the target analyte concentration can be determined by comparing the color on the T well with the color chart diagram.

Throughout the whole operation process, no external instrument nor electricity is required, enabling the proposed detector to be used for the detection of infectious disease in remote settings. Notably, the drying process and the subsequent washing step may compromise the performance of the antibodies as shown in Fig. S6, however, these procedures also can neglect the cost of refrigerated transport and improve the portability of the 3D-tPADs, which enable the on-site detection of the target analyte and extend the clinical application in resource-limited settings. In addition, the selectivity of the 3D-tPADs can be optimized by choosing the most prominent recognition receptors.29

3.2. Optimization of the washing and color development time of the 3D-tPADs-based ELISA test

For the proposed 3D-tPADs-based ELISA test, the washing buffer is added to rinse the dried detection antibody on the conjugate wells onto the detection wells, enabling antigen–antibody binding. However, after binding and forming the antigen–antibody complex, unbonded antibodies remain in the washing buffer. Therefore, the washing buffer needs to be completely removed since the residual antibodies will react with the TMB added in the next step and cause false positive or inaccurate ELISA detection results on the well. To remove this washing buffer, we included a stack of absorbent pads beneath the detection wells to soak up the excess solution, which generally required 4 min to completely eliminate the buffer from the detection wells.

In addition to optimizing the washing step, the color development time is also crucial for achieving accurate ELISA results.30 For the proposed 3D-tPADs-based ELISA test, the detection antibody modified with HRP is dried on the conjugate wells. After adding washing buffer to the conjugate wells, the HRP-detection antibody will be washed into the detection wells and later react with TMB solution to develop an intense blue color. The intensity of the color on the detection wells can be quantified using the blue to red color ratio (B/R) according to our previous findings.30,31 As we increase the color development time from 1 to 9 min, the B/R ratio increases, as shown in Fig. 2(a), with the highest B/R ratio achieved when the development time was 9 min. This condition produced a distinctive color difference between positive and negative trials, as shown in Fig. 2(a).

image file: d1lc00079a-f2.tif
Fig. 2 (a) The relationship between the color development time and the blue to red (B/R) ratio of the detection wells on the 3D-tPADs. (b) The HIV-1 p24 concentration as a function of the B/R ratio of the detection wells on the 3D-tPADs when the color development times were 1–9 min.

Additionally, an optimal color development time should result in a linear relationship between the B/R ratio and antigen concentrations in order to quantify the antigen in the test samples. Toward this aim, we performed a series of 3D-tPADs-based ELISA tests using antigen concentrations of 0 to 30 ng mL−1 and analyzed the resulting B/R ratio. We expected this ratio to linearly increase with the antigen concentration. However, with the color development time of 7 and 9 min, the B/R ratio derived from the antigen concentration of 3 ng mL−1 was higher than the one of 10 ng mL−1, which contrasts with the trend between the B/R ratio and the antigen concentrations of 0–1 ng mL−1. These contradictory results may be due to the saturation of TMB colorimetric assay, as shown in Fig. 2(b). Therefore, we chose 5 min as the optimal color development time as it results in good linearity between the B/R ratio and the antigen concentrations (Fig. 2(b)) as well as a relatively large B/R difference between the positive and negative results, as shown in Fig. 2(a).

3.3. Development of the electricity-free “timer” on the proposed 3D-tPADs

To indicate the optimal washing and color development times in a simple and automated manner, we integrated a timer function in the proposed 3D-tPADs. The “timer” starts when the washing buffer is added to the detection wells, subsequently flowing to the absorbent pads, then the dye wells, and finally to the indication wells. During this process, the washing buffer dissolves the dried green dye on the dye wells and colors the indication wells green, which serves as the signal (Fig. 3(a)). By carefully controlling the delivery time of the washing buffer, indication well 1 can be colored at the 4th minute and then indicator well 2 at the 9th minute through two different fluid paths as shown in Fig. S6 to indicate the appropriate times for the washing and color development steps, respectively.
image file: d1lc00079a-f3.tif
Fig. 3 (a) The schematic of the washing buffer flowing through the hydrophilic wells and channels on the detection pad, absorption pad, and indication pad. (b) The relationship between the delivery time and fluid path volume. (c) The delivery time as a function of the sucrose mass deposited on the absorption pad of the 3D-tPADs, and the corresponding schematic of the washing buffer flowing through a channel on the 3D-tPADs coated with dried sucrose, in which the viscosity of the washing buffer increases. (d) The delivery time as a function of pullulan mass deposited on the absorption pad of the 3D-tPADs and the corresponding schematic of the washing buffer flowing through a channel on the 3D-tPADs coated with dried pullulan, in which the viscosity of the washing buffer increases.

To deliver the washing buffer at the appointed times, we adjusted the volume of the fluid path and the viscosity of the fluid. To adjust the fluid path volume, we first created a fluid path by stacking four layers of absorbent pads to connect the hydrophilic wells and channels on different pads, as shown in Fig. 3(a). The fluid path volume can be increased by either increasing the thickness or the area of the hydrophilic channels and wells on the absorbent pads. As the fluidic path volume increases from 36.8 to 63.6 mm3, the fluid delivery time extends from ∼76.6 s to ∼571.3 s, as shown in Fig. 3(b) and S7. As a result, the required delivery times of 4 and 9 minutes can be achieved when the volumes are 45.2 mm3 and 49.7 mm3, respectively, as shown in Fig. 3(b). However, by adopting these fluid path volumes (45.2 mm3 + 49.7 mm3) to indicate the delivery times of 4 and 9 minutes, we would undesirably increase the device size, as shown in Fig. S8, leading to an increased cost of whole detector and also compromising the portability.

Therefore, to increase the delivery time while minimizing the device size, we resorted to another approach, which is to increase the viscosity of the washing buffer. Specifically, we dried sucrose and pullulan, which are commonly used and inexpensive reagents in paper-based devices,17,31,32 on the hydrophilic wells and channels of the absorbent pads (schematically showed in Fig. 3(a)). While flowing through the designed flow path on the pads, the washing buffer will dissolve the dried sucrose and pullulan, leading to an increased viscosity of the washing buffer and ultimately delaying the washing buffer delivery time. The resulting delayed delivery time depends on the mass of the sucrose and pullulan (for detailed information see Tables S1 and S2). Specifically, as we increase the mass of sucrose from 0 to 15.2 mg on the pads, the delivery time increases from 76.6 to 228.3 s (Fig. 3(c)). The appointed 4 min washing time (240 s) can be achieved when 15.2 mg of sucrose is evenly divided and deposited on each absorbent pad. However, when we further extended the delivery time to 9 min by increasing the sucrose mass to 3.2 mg, we observed a large variation in the delivery time (Fig. 3(c)), compromising the accuracy of the timer. Therefore, to achieve the appointed 9 min (540 s) needed to indicate the optimal TMB color development time, we deposited pullulan instead of sucrose on the fluid path used for indicating the appointed 9 minutes and increased the pullulan mass from 0 to 2.9 mg, resulting in an increase of the delivery time from 76.6 to 498.3 s, as shown in Fig. 3(d). As a result, a delivery time of ∼9 min can be achieved when the pullulan mass is 2.9 mg. Consequently, the expected washing buffer delivery times of ∼4 and ∼9 minutes can be achieved to indicate the completion of the washing and color development steps, respectively, independent of external instrumentation or electricity (Video S1).

3.4. Detection of HIV-1 p24 on the 3D-tPDAs

To demonstrate the feasibility of the proposed origami-based 3D-tPADs design, we use the device to detect HIV. We chose the viral capsid protein HIV-1 p24 as the target antigen, since it has long been recognized as a virological biomarker in early HIV detection.28 Using the proposed sensor, we establish the standard curve of the HIV-1 p24 protein concentration, as shown in Fig. 4(a). Specifically, we performed the detection with PBS containing 0 to 30 ng mL−1 of HIV-1 p24 antigen and analyzed the resultant B/R values of the detection wells. We found the B/R values increase as we increase the HIV-1 p24 protein concentration. The dynamic linear range was from 0.03 ng mL−1 to 3 ng mL−1. The coefficient of determination (R2) was 0.98 with a limit of detection of 0.01 ng mL−1.
image file: d1lc00079a-f4.tif
Fig. 4 (a) Images of the 3D-tPADs ELISA results with HIV-1 p24 concentrations ranging from 0–30 ng mL−1, and the corresponding B/R ratios. (b) Images of the 3D-tPADs ELISA results with plasma spiked with 1–30 ng mL−1 of HIV-1 p24, and the resulting B/R ratios. (c) Semi-quantification of the HIV-p24 detection results using the color chart diagram.

To validate the feasibility of the proposed 3D-tPADs in detecting clinical samples, we further performed the tests with human plasma spiked with 0 ng mL−1 to 30 ng mL−1 of HIV-1 p24 protein. The resulting B/R values of the detection wells increased as we increased the HIV-1 p24 protein concentration in the plasma, as shown in Fig. 4(b). The linear detection range was from 0.03 ng mL−1 to 3 ng mL−1 with an R2 of 0.97 and the detection limit was 0.01 ng mL−1, which is comparable or superior to other related assays as shown in Table S3. Based on this linear relationship, an unknown HIV-1 p24 concentration can be quantified, verifying the ability of the 3D-tPADs design to process clinical samples.

The proposed 3D-tPADs is modified with CMC (see the Experimental section for details), which helps to reduce non-specific binding of random proteins to its surface, providing distinct advantages including low background noise and strong signal intensity.33 Benefiting from these advantages, positive HIV-1 p24 detection results can be distinguished from negative ones by the naked eye, as shown in Fig. 4(a and b). In addition, detection results based on different HIV-1 p24 concentrations can be easily differentiated by the naked eye due to the large B/R value differences between the images, as shown in Fig. 4(a and b). Taking advantage of the high color variance of the 3D-tPADs, we developed a result readout method that does not require electricity. Specifically, we developed a color chart diagram that shows the detection results of the 3D-tPADs with different HIV-1 p24 concentrations. After performing the detection on the 3D-tPADs, the HIV-1 p24 concentration can be semi-quantified by comparing the color change of the 3D-tPADs to the color chart diagram (Fig. 4(c)). The detection limit of this semi-quantification method is 0.03 ng mL−1, since the detection result based on this concentration can be distinguished from the negative result (0 ng mL−1) by the naked eye (Fig. 4(a and b)). Consequently, the HIV-1 p24 detection result can be achieved independent of electricity or external equipment, such as a smartphone or computer, as required in traditional 3D-tPADs analysis.

4. Conclusions

In this study, we develop a fully integrated infectious disease sensor that can detect disease independent of electricity or external instrumentation. The sensor consists of a 3D-tPADs that is designed according to the principle of origami. By folding and sliding the device tabs, we are able to control the reagent/sample flow through the different assay media pre-loaded in the paper substrate, allowing the multi-step ELISA test to be easily performed in the correct sequence, with little training necessary. To indicate the timing of the ELISA steps, we further developed a “timer” function, in which colored fluid simultaneously travels to the indicator pad of the 3D-tPADs at a controlled flow rate. We verified the feasibility of this timer-integrated 3D-tPADs by performing HIV-1 p24 detection, in which we observe an intense blue color when the samples contain HIV-1 p24 and a light blue color when they do not. In addition, plasma samples can be directly applied onto the 3D-tPADs for HIV-1 p24 detection. Furthermore, the HIV-1 p24 concentration can be semi-quantified by comparing the color change of the 3D-tPADs to the sensor's pre-prepared color chart, allowing the result to be measured without the necessity for image processing as in traditional 3D-tPADs, and with a detection limit as low as 0.03 ng mL−1. Throughout the whole detection process, no electricity nor external instrumentation is required, enabling the proposed sensor to be used in remote settings for on-site infectious disease diagnosis. To further extend the application of the proposed 3D-tPADs in point-of-care settings while maintaining the immunoassay performance, stabilizers such as sugars, amino acids and salts can be added onto the device to protect the pre-dried antibodies from denaturation, and also the device can be stored in a sealed aluminium foil bag to protect from light, gas and moisture.30,34–37 In addition, other types of capture/detector agents such as aptamer and DNA can be used to replace the currently used p24 antibody to further extend to the range of detectable disease biomarkers. Notably, to ensure proper selectivity of the 3D-tPAD, the antibodies or agents with fast kinetics and excellent binding affinity to the target of interest should be used.29

Author contributions

C. F. C. initiated the research; C. A. C., H. Y., C. W. C., Y. S. C., and W. H. S. performed the experiments; all authors participated in analyzing the data and writing the paper.

Conflicts of interest

There are no conflicts to declare.


This research was supported by the Ministry of Science and Technology, Taiwan (109-2223-E-002-004-MY3, 110-2923-E-002-004-MY3, and 108-2923-E-002-004-MY2), and the Higher Education Sprout Program at National Taiwan University (110L891504).


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Electronic supplementary information (ESI) available. See DOI: 10.1039/d1lc00079a
Equal contribution.

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