Stefano Begoloa, Feng Shenb and Rustem F. Ismagilov*a
aDivision of Chemistry and Chemical Engineering, California Institute of Technology, 1200 East California Boulevard, Pasadena, CA 91125, USA. E-mail: rustem.admin@caltech.edu
bSlipChip Corp., 129 N. Hill Ave., Pasadena, CA 91106, USA
First published on 17th September 2013
This paper describes a microfluidic device for dry preservation of biological specimens at room temperature that incorporates chemical stabilization matrices. Long-term stabilization of samples is crucial for remote medical analysis, biosurveillance, and archiving, but the current paradigm for transporting remotely obtained samples relies on the costly “cold chain” to preserve analytes within biospecimens. We propose an alternative approach that involves the use of microfluidics to preserve samples in the dry state with stabilization matrices, developed by others, that are based on self-preservation chemistries found in nature. We describe a SlipChip-based device that allows minimally trained users to preserve samples with the three simple steps of placing a sample at an inlet, closing a lid, and slipping one layer of the device. The device fills automatically, and a pre-loaded desiccant dries the samples. Later, specimens can be rehydrated and recovered for analysis in a laboratory. This device is portable, compact, and self-contained, so it can be transported and operated by untrained users even in limited-resource settings. Features such as dead-end and sequential filling, combined with a “pumping lid” mechanism, enable precise quantification of the original sample’s volume while avoiding overfilling. In addition, we demonstrated that the device can be integrated with a plasma filtration module, and we validated device operations and capabilities by testing the stability of purified RNA solutions. These features and the modularity of this platform (which facilitates integration and simplifies operation) would be applicable to other microfluidic devices beyond this application. We envision that as the field of stabilization matrices develops, microfluidic devices will be useful for cost-effectively facilitating remote analysis and biosurveillance while also opening new opportunities for diagnostics, drug development, and other medical fields.
Generally, analytes are kept stable using the “cold chain,” in which samples are transported in dry ice and stored in freezers. The cost and complexity of low-temperature stabilization limit the applications of biosurveillance and follow-up tests.7 Furthermore, freezers that house specimens require electricity, putting samples at risk of destruction in the event of power failure.8,9 And while the cold chain addresses the question of storing specimens, it does not account for other needs of remote analysis. For instance, many biospecimens must be specially prepared at the moment of collection in order to be studied further. In the case of blood biomarkers, analysis is typically performed on serum or cell-free plasma,10 which must be obtained by trained personnel using specific equipment, such as a centrifuge.
Dried blood spots (DBS) have been used as one alternative to the cold chain. This method involves drying whole blood on filter paper to stabilize the sample. Recovery is performed by removing a portion of the blood spot with a hole punch and using a combination of solvents and buffers to elute the sample from the resulting punched-out disc. This method does not require sample preparation or specialized equipment at the moment of sample collection, and has been used for large-scale neonatal screening and biosurveillance.11 However, the use of DBS for quantitative analyses presents some challenges and limitations, as results are dependent on the hematocrit and on where the punch is taken on the paper,12 and samples can be contaminated by open-air exposure or cross-contaminated by the biopsy punch tool. Furthermore, long drying times and external conditions such as humidity may compromise the stability of analytes: for this reason, most protocols require freezing of DBS for long-term storage.13,14 Finally, serum or cell-free plasma, rather than whole blood, are often the matrices of choice for clinical tests; to implement DBS, specific elution procedures need to be optimized for purifying analytes and avoiding interference from residual components in blood. Although dried plasma spots can address some limitations of DBS, they require trained personnel to perform plasma separation.
Chemical stabilization matrices offer an alternative to the cold chain and DBS, but until now, they have not been compatible with devices for resource-limited settings. A number of matrices have been developed, many of which rely on natural chemistries used by living organisms for self-preservation.15 Several commercially available stabilization matrices (Biomatrica, GenTegra, etc.) preserve samples in the dry state. Drying inhibits analyte degradation linked to enzymatic activity and slows down reactions that are activated by light or heat.16,17 These matrices do not necessarily solve the problem of sample preparation, but they do allow long-term storage of DNA and RNA at room temperature. Current protocols for using these stabilization matrices are suitable for laboratory use, but drying the samples typically requires vacuum conditions or external devices (e.g., lyophilizer, pumps, vacuum concentrators) to control humidity, as well as experienced personnel to process the sample.18,19 Portable devices have been proposed,20 but these usually involve electricity and long drying times (from a few hours to overnight). Alternatively, drying can be done without the use of external equipment by exposing the sample to the environment, but airborne contaminants may come into contact with the sample and drying times are dependent on external conditions such as humidity. Moreover, leaving the tubes or titer plates open during drying may lead to cross-contamination between different samples: to solve this problem, one needs to use porous seals (which increase drying times) or a dedicated device20 for preventing aerosol-driven cross-contamination.
An approach that offers reliable preservation, avoids contamination, allows quantitative measurements, is compatible with sample preparation, and can be executed by minimally trained users is needed. Herein, we describe a device that meets these requirements and can be used with commercially available sample preservation matrices.
Slipping activates drying by putting the sample in vapor contact with the molecular sieves. A polycarbonate porous membrane embedded in P2 prevents contact between sample and desiccant at this stage. Drying is selectively activated: P2 and P3 both have a series of channels that do not overlap in the “Loading” and “Recovery” positions. Only the “Drying” position aligns these channels, thus creating the vapor contact that drives the drying. Drying dynamics and timescale were characterized by taking videos or photographs of the channels using a camcorder or a camera.
The recovered solution can be handled using the pipettor and analyzed with conventional techniques. We tested the recovery efficiency using water containing green food dye, Phosphate buffered saline (PBS) with Alexa Fluor 488 (Sigma Aldrich), or CdTe/Cds core/shell Quantum Dots (QD). QDs were synthesized following a published method,22 with core size between 2.5 and 3 nm and stock concentration on the order of 2.6 μM.
Samples were diluted to the desired concentration (80 ng μL−1 for control RNA, 375 copies per microliter for HIV-1 RNA) and mixed with a chemical stabilization matrix for RNA (RNAStable, purchased from Biomatrica, Inc.).
Several aliquots of these solutions were injected into the device and dried for stabilization. The devices were then stored at 50 °C for up to 5 weeks. In parallel, several aliquots of the same solution were used as controls. Controls included aliquots stored in a −80 °C freezer and aliquots stored in the liquid state at 50 °C in microcentrifuge tubes. Each storage condition was performed in triplicate. At each time point, the sample was recovered from the devices using the rehydration and re-collection procedures, and three washes were performed for each well. All solutions were then diluted so that the total volume of each aliquot was 100 μL.
Fig. 1 An overview of the operations and capabilities of the device. (A) A photograph representing the first step, in which an untrained user places the collected sample (here, a green aqueous solution) at the device inlet. (B) A photograph of the device after the lid has been placed (left), which activates the automated filling process that enables precise volume quantification (right). Sample preparation, such as plasma filtration, is integrated in this step. (C) A zoomed-in photograph of the device after the user has performed the third step: a slip that disconnects the aliquots and activates drying. (D) A photograph showing the device after complete drying. (E) A photograph of an agarose gel (inverted intensity) showing clear bands and no degradation, indicating that samples stored in the device remain stable without the need for refrigeration. (F) A photograph of the device with one rehydrated well (green). Aliquots can be re-collected independently, so part of the sample may be kept dried for further storage. (G) A graph showing a standard qPCR profile; recovered samples can be analyzed using traditional methods and instruments. Scale bars are 5 mm in all photographs. The loading process is also shown in Video S1, ESI.† |
The sample, which stays fully enclosed in the device, remains stable and in the dry state during transport and/or storage (Fig. 1E). At the desired time, a trained user can perform a second slip of the device with a special “slipping tool” and rehydrate one or more of the aliquots by injecting water into the wells (Fig. 1F) as detailed below. These rehydrated aliquots can then be re-collected using a standard pipettor and are ready for further analysis or purification (Fig. 1G). If not all aliquots are needed for analysis, aliquots that are not rehydrated can be stored long-term (e.g., for archival purposes). Although the devices described in this paper are prototypes, they were designed in consultation with manufacturers to ensure compatibility with mass production.
Fig. 2 Automated loading with precise volume quantification. (A) A photograph (top) and a schematic drawing (bottom) of an empty device, with a sample (green solution) placed at the inlet, with a flexible o-ring surrounding the inlet. Lines have been added to the photograph to illustrate the geometry (dashed lines outline ducts in P1 (one of the device’s three major parts, or subassemblies), and solid, light gray lines outline wells in P2). A light gray circle was used to indicate the via hole used for dead-end filling, described in more detail in the ESI† (Fig. S5). (B) Photograph and schematic demonstrating that as the lid is placed on the device, it forms a tight seal with the o-ring, creating pressure and initiating loading. Wells are loaded sequentially (following the path indicated by arrows) so that each well is completely filled before the solution enters the next well. (C1) A photograph of a fully loaded device, in which the sample volume (55 μL in this case) is larger than the total device capacity (50 μL). All wells are filled, and loading has stopped automatically. (C2) A photograph of a partially loaded device, in which the sample volume (25 μL in this case) is lower than the total device capacity. Loading stops automatically as soon as air enters the first well (“venting well”), allowing quantitative measurements to be obtained by counting full wells. |
The second design feature crucial for loading involves a combination of dead-end and sequential filling that together allow precise volume quantification. Dead-end filling is a strategy previously applied for the loading of SlipChip devices,24 in which the fluid (e.g., oil or air) originally present in the wells or channels of the device is evacuated through the gap between the device layers as the sample is entering the wells or channels. In the cases previously reported, the “sealing pressure” (defined as the maximum pressure that can be used for loading the device without the sample entering the gap) is inversely proportional to the gap between the device layers (see model described in ref. 24).
In the experiments described in this work, the air originally present in the wells of the device was pushed through the membrane pores and evacuated through a via hole placed in the proximity of the last well. A schematic description of this mechanism is provided in the ESI† (Fig. S5). We emphasize that the liquid sample is injected into the device wells and channels only, and does not penetrate the porous membrane that serves as the bottom of the device’s wells. This configuration increases the range of loading pressures, as the sealing pressure24 in this case is controlled by the size of the pores in the membrane. As an example, when loading blood plasma (surface tension 50 mN m−1) into a well filled with air and using a membrane with a pore size of 0.45 μm, the sealing pressure for the membrane is ~2 bar (roughly 4 times more than the sealing pressure for a 1 μm gap between plates). This value is significantly higher than the pressures generated by the pumping lid, and we did not observe any permeation of liquid through the porous membrane. Choosing a membrane with an even smaller pore size would further increase the sealing pressure, making the device resistant to even higher loading pressures.
Filling stops automatically when all the wells are filled with the sample, and any extra volume placed on the device inlet is not injected into the device, providing an effective control on the maximum loaded volume. With sequential filling (Fig. 2B), each well fills completely before the liquid reaches the following well. If the input volume is lower than the device capacity, loading stops automatically as soon as the entire sample has been injected (Fig. 2C). Any extra air that follows the end of the liquid sample slug can be vented through an extra “venting well” in the storage layer to stop filling. The original volume of sample can then be quantified by counting the number of full wells or by taking a picture of the device after filling and analyzing it using ImageJ Software to approximate the fraction of each well containing the solution. Repeated loading experiments demonstrated that filling is consistent with these predictions (Fig. 3).
Fig. 3 A plot comparing injected volume as measured by the device with the original sample volume. The relation is linear below 50 μL, which is the maximum device capacity. |
These design features obviate the need for users to precisely control pressure and input volume, both of which are challenging operations without proper training and equipment. Knowledge of the input volume is critical for calculating concentrations of analytes in the biofluid being preserved and then analyzed. Moreover, we demonstrated that a wide range of solutions can be loaded using this approach (see Table S1 and Video S1 in the ESI†). The large flexibility in pressures, ensured by the modified dead-end filling mechanism described above, made it possible to rapidly drive fluids with the viscosity of water (with velocities up to 10 μL s−1) through the device. This loading approach was also suitable for loading samples with a wide range of properties, including viscosities up to 110 mPa s (85% w/w of glycerol in water) and surface tensions down to 7 mN m−1 (0.4 mM BSA). For this BSA solution, the calculated sealing pressure was 0.31 bar.
In this work, the matrix was pre-mixed with the sample prior to injection into the device. The device has been designed to be compatible with a variety of matrices, even those not yet developed. The preferred method of incorporating a specific preservation matrix into the device would depend on the properties of the particular matrix and its dissolution kinetics.
After the user has performed the slip, the drying process in each well begins with the nucleation of an air bubble that grows as evaporation occurs (Fig. 4). We relied on a well-understood idea25–28 that by reducing length scales over which evaporation takes place, vapor transfer can be accelerated. This process is rapid also because evaporation takes place through the porous membrane that serves as the bottom layer of the wells. The surface to volume ratio in this case was 3.3 mm2 for each μL. Drying times for the different solutions tested were typically on the order of 30–45 min for a total volume of 50 μL (Fig. 4E) (see Fig. S6 and Video S2 in the ESI† for drying dynamics). As a comparison, the same volume in a microcentrifuge tube required up to 12 hours to be evaporated in a laboratory environment. These timescales were suitable for the storage experiments shown in this paper, but this framework offers a wide range of drying times that can be adjusted by changing device parameters such as the volume of wells, surface-to-volume ratio, type of desiccant, and membrane properties (such as pore size, fractional open area of the membrane, and membrane thickness). After drying is complete, the presence of sample in each well can be confirmed by the presence of solid residuals (Fig. 4C and D), which can be used as a strategy to detect empty wells in the event of partial filling.
Fig. 4 An overview of slip-initiated drying. (A) A photograph (top) and schematic drawing (bottom) showing the device after loading. In this case, the device was loaded with a pipettor instead of using the “pumping lid” to ensure unobstructed visualization of the sample wells. (B) A photograph and a schematic drawing of the device once it has been slipped. The slip disconnects wells and allows vapor contact between the sample and the desiccant (black). Evaporation starts by nucleation of an air bubble in each well. (C) A photograph and a schematic drawing showing drying in progress. The bubbles grow due to progressive loss of water caused by evaporation. (D) A photograph and a schematic drawing of the device once drying is complete and the sample is stabilized (green sample residuals are visible in the wells). (E) A plot showing the fraction of the wells containing liquid over time for a representative device. See ESI† (Fig. S6) for full schematic drawings and operation of the device. |
Fig. 5 An overview of sample rehydration and recovery. (A) A photograph (top) and a schematic drawing (bottom) of the device containing dry sample after 12 hours, along with a drawing of a separate “slipping tool” used for slipping to the recovery position. (B) A photograph and a schematic drawing showing a device that has been slipped to the recovery position using the special slipping tool. (C) A photograph and a schematic drawing of the device after selective rehydration of a single well with a pipettor. (D) A photograph and a schematic drawing showing selective recovery of the sample from a single rehydrated well (other aliquots remain dry for further storage). Full schematics and operation of the device in A–D are available in the ESI† (Fig. S7). (E) A graph showing quantitative recovery of quantum dot solutions after drying has taken place in the device. (F) A graph showing quantitative recovery of Alexa Fluor 488 solutions after drying has taken place in the device, compared to the original solution (“control”). In E and F, brackets indicate p values, the null hypothesis being that the two concentrations were the same in each case. Error bars represent the 95% confidence interval (n = 3 for each condition). See Fig. S8 and S9† for calibration curves, Tables S2 and S3† for statistical calculations, and Video S3† for demonstration of rehydration and recovery in the ESI†. |
After rehydration, the sample can be retrieved from each well by aspirating with a standard pipettor (Fig. 5D). Using this procedure, we re-collected liquid solutions with the same concentration as the injected sample, demonstrating that quantitative measurements can be obtained from stored aliquots (Fig. 5E and F). We evaluated the reliability of recovery by injecting solutions containing either fluorescent molecules (Alexa Fluor 488) or nanoparticles (CdTe/Cds quantum dots) (Fig. 5E and F). After drying and rehydration, the solutions were re-collected from the wells of the device. This method allowed quantitative measurements: solutions with different concentrations were reliably distinguished, showing clear difference between solutions with a 2-fold difference in dilution (all p values were 5 × 10−5 or less, the null hypothesis being that the two concentrations were the same in each case). No more than ±10% difference was observed when comparing the signal from the rehydrated aliquots with the starting stock solution (see Tables S2 and S3 in the ESI†). If needed, liquid residuals in the channel can be washed out by injecting more water into the well and re-collecting the solution; the washing step can be repeated additional times if necessary.
Since each well is individually retrievable, a trained user can rehydrate one or more wells and perform either partial or total recovery of the original sample. If only a portion of the total number of wells is recovered, the device can be slipped back to the drying position for further storage.
Fig. 6 Integration of the sample preservation device with a plasma filtration module. (A) Schematic drawings that show the components of the plasma filtration module (top) and the process for obtaining cell-free plasma with this module (bottom). The pumping lid is used both to obtain plasma and to load the device. (B) Two photographs of the device after the lid has been closed and plasma has been successfully obtained from whole blood and loaded into the wells. Whole blood was spiked with 20 mM of Alexa Fluor 488 to aid visualization of the separated plasma. Left: Bright-field image, showing the red and white blood cells (residuals) at the device inlet and the plasma loaded in the wells. Right: Fluorescence image of the device (for improved visualization of the plasma within the wells). (C) A graph comparing quantification results of HIV-1 RNA from plasma extraction with a traditional centrifuge (left) and extraction using the device’s plasma filtration module (right). p Values are in the same range for both techniques: *** indicates a p value of less than 0.001, and n.s. indicates a p value higher than 0.05 (no significant difference) (see Table S4 in the ESI† for full details of statistical analysis). The null hypothesis for p values was that both concentrations were equivalent. Error bars represent the 95% confidence interval (n = 3 for all experiments, except for the centrifugation experiment at the lower concentration, for which n = 5). |
Using this filtration approach, we successfully separated plasma from whole blood that was spiked with HIV-1 viral particles. We were able to purify HIV-1 RNA from these plasma samples. Compared to plasma controls separated by centrifugation, the recovery rate of HIV-1 RNA after plasma filtration with this device was always more than 60%. We used different concentrations of HIV-1 particles and found that the p values were comparable for the control samples and the device (1.2 × 10−4 for the centrifuge and 1.6 × 10−4 for the device) (Fig. 6C) (see Table S4 in the ESI† for p value calculations).
Since there is currently no chemical matrix available for stabilization of HIV-1 RNA in plasma, we did not test the stability of preserved RNA in plasma. However, we verified that the device can successfully dry normal plasma. We demonstrated that injected plasma can be dried down, stored, and re-collected as described earlier (see Fig. S10 in the ESI† for details). Rehydration times for plasma were longer (on the order of 15–20 min) than for the previous experiments, so we typically injected the water back and forth with a pipettor to speed up the dissolution. One critical parameter is the speed at which the plasma samples are dried. Since, in the dry state, blood plasma produces more solid residuals than purified solutions, there is a risk that the wells may get clogged when the sample dries. The device’s fast drying times avoided this problem by distributing residuals across the walls of the wells, allowing successful rehydration of all wells (Fig. S10 in the ESI†). In contrast, when we tested the drying of plasma through the porous membrane using the P2 subassembly alone (not integrated with the device) and no desiccant to drive the drying process, times for drying were on the order of 1 to 2 hours, and the solid residuals tended to accumulate in one spot in the wells and block the fluidic path for rehydration.
To test general RNA stability, we used a mixture of control RNA 250 (Ambion) and a stabilization matrix (RNAStable, Biomatrica). Electrophoresis experiments performed with an Agilent 2100 Bioanalyzer showed that the 80 ng μL−1 of control RNA aliquots stored dry in the device for 4 days at 50 °C did not show any detectable degradation, indistinguishable from the controls stored in a freezer at −80 °C (see Table S5 and Fig. S11 in the ESI† for full details, evaluated semi-quantitatively using an approach similar to the one described previously).31,32 As a comparison, degradation of control RNA was obvious in aliquots stored in the liquid state at 50 °C, evidenced by the presence of short products that formed a smear pattern (Fig. 7A).
Fig. 7 Validation of the device using control RNA and HIV-1 RNA. (A) Densitometry plot showing electrophoresis results from control RNA (80 ng μL−1) mixed with a stabilization matrix (RNAStable, Biomatrica) and stored for four days under different conditions. (B) A graph showing the results of quantitative analysis of purified HIV-1 samples performed with RT-qPCR. Error bars represent the 95% confidence interval (n = 3). Stars are used to indicate p values: * if the p value is less than 0.05, ** if less than 0.01, and *** if less than 0.001. See Table S7 in the ESI† for calculations. |
Next, we showed that the device also allowed stabilization and quantitative recovery of HIV-1 RNA. RNA was purified from inactivated HIV-1 viral particles (see Experimental). Aliquots (10 μL) containing ~3750 copies of RNA and mixed with the preservation matrix were stored in one of three conditions: (i) frozen at −80 °C, (ii) in the device in the dry state at 50 °C, or (iii) in the liquid state at 50 °C in microcentrifuge tubes. Experiments at elevated temperatures are typically used to evaluate long-term stability of samples in a reasonable amount of experimental time; furthermore, such experiments reflect temperature fluctuations to which samples may be exposed during shipping when temperature is not controlled. Since this RNA was at a low concentration (375 copies μL−1), RT-qPCR was used to evaluate stability and recovery rate. Quantification cycles (Cq) were measured for each storing condition and were compared with controls (aliquots stored at −80 °C) (Fig. S12 and S13 and Table S6†). Liquid samples stored at 50 °C showed significant degradation after as few as 7 days (p value = 0.04), and this degradation became more prominent over time (p value after 35 days was 6.1 × 10−8) (see Fig. 7B and Table S7 in the ESI†). When comparing the frozen controls and those samples stored in the device at 50 °C, we observed a minor difference in Cq values (0.5 cycles on average, n = 18), but no substantial change (always less than 0.3 cycles) in Cq was observed over a 5-week period. Stability of the RNA at 50 °C over a 5-week period in this “accelerated aging” test17,33 implies that the RNA may be stable at room temperature in the device for as long as 8 months.17,33
Now that we have demonstrated and validated the features of the device such as pumping, modular design, integration of plasma filtration, and aliquoting, the next step is to develop and test new matrices to stabilize desired targets, such as HIV-1 RNA in plasma. We emphasize that in this work, we did not create new chemical stabilization matrices and used only those commercially available. The range of applications of this device will expand as additional matrices are being developed in this active field of research. We envision that the device will be used for remote analysis, allowing quantitative tests such as quantifying viral load or genotyping for viral infection (of HIV, HCV, etc.). While we focused on preservation and recovery of nucleic acids, we have demonstrated that this approach is also compatible with recovery of small molecules and particles. Recovery of small molecules is attractive for studying pharmacokinetics in clinical trials of drug candidates.10,11 Preservation of proteins and live organisms using this approach is of active interest to us as well.
Examples of other potential applications include the testing of different analytes from a single sample (multiplexing) by using more than one stabilization matrix, as well as multi-patient testing achieved by the storage of more than one sample in the same device. In addition, sample stabilization during transport could also facilitate large-scale, remote clinical trials by allowing subjects to collect samples at home in order to minimize the costs of traveling to a centralized facility, sample collection, and analysis. Another potential application of analyte stabilization is the ability to track the clinical history of a patient by preserving samples collected regularly over a long period of time. This would dramatically increase the amount of available information for personalized medicine, allowing one to trace the temporal evolution of disease and biomarkers not only after the diagnosis, but potentially over a patient’s lifetime. This approach is currently unfeasible due to the high costs of cold-chain stabilization and biobanking, but it would be enabled by further developments of bio-preservation reagents and of integrated, simple-to-use microfluidic devices of the type described here.
Footnote |
† Electronic supplementary information (ESI) available. See DOI: 10.1039/c3lc50747e |
This journal is © The Royal Society of Chemistry 2013 |