Fabrication of Tapered Microtube Arrays and Their Application as a

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Fabrication of Tapered Microtube Arrays and Their Application as a Microalgal Injection Platform Andrew R. Durney,† Leah C. Frenette,‡ Elizabeth C. Hodvedt,† Todd D. Krauss,‡ and Hitomi Mukaibo*,† †

Department of Chemical Engineering, and ‡Department of Chemistry, University of Rochester, Rochester, New York 14627, United States S Supporting Information *

ABSTRACT: A template-synthesis method that enables fabrication of tapered microtube arrays is reported. Tracketched poly(ethylene terephthalate) membranes are used as the template, with closed-tipped conical pores having length and base diameter of 6.27 ± 0.28 and 1.21 ± 0.05 μm, respectively. A conductive layer of Pt is deposited by atomic layer deposition (ALD) to enable the successive electrodeposition of Ni. By decreasing the Pt precursor pulse duration from 10 to 1 s during the ALD step, the heights of the microtubes are controlled from the maximal full length (∼6 μm) to only a fraction (1−2 μm) of the template pore. Using a pulsed-current electrodeposition (PCD) method, a smooth and uniform Ni deposit is achieved with a thickness that can be controlled as a function of the PCD cycle. The microtubes’ lumen is confirmed to stay open even after 2000 cycles of Ni PCD. A potential application of the prepared array as a microinjection platform is demonstrated via successful injection of 10 nm sized CdZnS/ZnS core/shell quantum dots into Chlamydomonas reinhardtii microalgae cells with intact cell walls. The direct delivery method demonstrated in this paper offers novel opportunities for extending the growing interest in array-based microinjection platform to microalgal systems. KEYWORDS: bionanotechnology, characterization tools, microstructures, thin films, quantum dots

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INTRODUCTION The applications of vertically protruding tapered microtube arrays (TMAs) are immensely diverse: they range from sensors,1−4 plasmonics,5−8 and solar energy harvesting9,10 to battery electrodes11,12 and transdermal drug delivery.13,14 While numerous lithography-based technologies are developed to fabricate these three-dimensional structures,15−20 these methods have become increasingly sophisticated and expensive as the required microtube dimensions become smaller.21,22 Alternatively, a membrane-based template synthesis method23 has been applied as an inexpensive, generic platform for preparing arrays of submicrometer-sized microtubes with various materials and dimensions,24−26 including TMAs.27 In this method, a thin, nanoporous membrane (such as anodized aluminum oxide or track-etched polymer film) is used as a sacrificial template. The array is obtained by depositing a thin layer of the desired material into the template pores and subsequently removing the template membrane. The outer geometry of each microtube is defined by the geometry of the template pore, and the inner lumen is formed by limiting the thickness of the deposited layer.26,28−31 In this manner microtube arrays with tapered walls can easily be prepared by depositing a thin layer of the desired material to a cone-shaped template pore. However, conventional approaches that deposit the desired material on the entire pore wall surface can only yield open© XXXX American Chemical Society

tipped TMAs from pores that penetrate the entire thickness of the membrane (i.e., pores that are open through to both membrane faces). This is an issue for fabricating an array of short TMAs because the template membrane becomes increasingly fragile with decreasing membrane thickness. Recently, Li et al. have shown that electrodeposition can be used to prepare open-tipped nanocone arrays from closedended, conical nanopores that are embedded within a thick, mechanically robust template membrane.27 This result was achieved due to a geometric shadowing effect that limits the penetration depth of the sputtered Au used as a cathode for the electrodeposition.32 However, the bases of the nanocones were fully clogged due to high current density at the pore opening during electrodeposition, and furthermore, the shadowing effect prevents this approach from yielding nanocones with high aspect ratio. In this paper, we describe a template-synthesis method that combines atomic layer deposition (ALD) and pulsed-current electrodeposition (PCD) to yield an array of tapered microtubes with channels that are open on both ends and with heights that are not limited by the shadowing effect of the template pores. We also demonstrate the potential application Received: September 1, 2016 Accepted: November 23, 2016 Published: November 23, 2016 A

DOI: 10.1021/acsami.6b11062 ACS Appl. Mater. Interfaces XXXX, XXX, XXX−XXX

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ACS Applied Materials & Interfaces

Scheme 1. Cross-Sectional Illustrations of (a) Pt, Au, and Ni Layers Deposited on TE-PET Template Pores, (b) Fully Exposed TMA on Cu Foil Substrate, (c) Partially Exposed TMA with Residual Template Membrane, and (d) a Pierced C. reinhardtii Cell on a TMA and Delivery of QDs

of this TMA as a microinjection platform for the widely studied microalgae Chlamydomonas reinhardtii.33,34 Microalgae are a diverse group of aquatic, unicellular, photosynthetic organisms that are studied intensively for their ability to produce highvalue-added products such as biofuels, recombinant proteins, pharmaceuticals, and fine chemicals. However, the hard cell wall has proven to be a critical barrier that hinders the advancement of microalgal technology.35 Our study offers novel opportunity to bypass the cell wall barrier and to extend the growing microtube-array-based injection technology from the widely studied mammalian systems24,36−41 to the much smaller microalgal systems.

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prepared using a Au electroless plating method to replicate original template pores with high fidelity.31,50 The top-down view of a single Au replica in the Figure 1b inset shows that the nanocone tips are sharp and fully closed. This is expected from the fully closed tip of the template pore (Figure 1a). From these replicas, the length and the base diameter of the template pores were measured to be 6.27 ± 0.28 and 1.21 ± 0.05 μm, respectively, with tip diameters less than 100 nm. To create TMAs using this template, the penetration depth of the conductive seed layer into the template pore needs to be restricted to prevent Ni electrodeposition at the tips of the closed template. This concept is demonstrated in Figure 1c, where sputtering is used to coat the TE-PET template surface with a metallic layer of Au. Since sputtering is susceptible to shadowing effects on high-aspect-ratio structures, penetration of the Au layer into the submicrometer pores is limited.27,32 As a result, successive Ni electrodeposition occurs only at the vicinity of the pore entrance, resulting in structures that are 2.44 ± 0.39 μm talla third of the length of the template pore. The base diameter was 1.08 ± 0.03 μm, which is consistent with the base opening of the template pore. The tip diameter is 0.66 ± 0.04 μm, which is consistent with the diameter of the template pore at ∼1/3 depth. The thickness of the Ni deposit is controlled to yield hollow structures, as shown in the inset of Scheme 1c. Effect of Pt ALD on Microtube Height. In order to fabricate taller TMAs that are not limited by the template’s shadowing effect, Pt ALD was used to deposit a conductive seed layer with controllable penetration depth into the pores.42−47 The Pt precursor pulse duration was controlled to tune the penetration depth of Pt precursor into the template pore. The ALD protocol was developed based on a previously reported low-temperature recipe to prevent melting of the polymeric template membrane.51 An additional 30 nm of sputtered Au film was deposited on top of the 15 nm thick Pt film to improve conductivity and enable sufficient Ni electrodeposition (Scheme 1a). The TMA shown in Figure 2a was prepared with a short (1 s) Pt precursor pulse duration. These short-pulse microtubes have heights of 2.68 ± 1.32 μm and base and tip outer

RESULTS AND DISCUSSION

Approach. Track-etched poly(ethylene terephthalate) (TEPET) membranes with conical pores were prepared following a method described previously.33 A Pt conductive seed layer was deposited onto the surface of the insulating TE-PET prior to Ni PCD. The penetration depth of the Pt layer into the template pore was controlled following previous ALD approaches (Scheme 1a).42−47 A thin layer of sputtered Au was added to enhance conductivity. As described below, the prepared conical pores were much deeper than the penetration depth of the sputtered Au. PCD was chosen specifically to achieve smooth deposits with precisely controlled thicknesses.48,49 The TMAs were exposed by removing the TE-PET and then characterized using field-emission scanning electron microscopy (FE-SEM), focused ion beam (FIB) milling, and spectroscopy methods (Scheme 1b). A thin TE-PET layer was left on the surface of the TMAs used in ion-current resistance measurements (Scheme 1c) and cell piercing experiments (Scheme 1d) to provide stability during handling. Fabrication of TMAs. Figure 1a shows the cross section of a template pore, revealed using a dual-beam focused ion beam scanning electron microscope (FIB-SEM). Together with the top-down view into the pore (Figure 1a, inset), the conical shape with a wide base opening that narrows down to a sharp closed tip is confirmed. The uniformity across the multiple pores is better visualized in Figure 1b, showing a representative side view of multiple pore replicas. These replicas were B


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Figure 2. FE-SEM images of TMAs prepared with Pt ALD using (a) 1, (b) 5, or (c) 10 s Pt precursor pulse duration followed by 250 cycles of Ni PCD. Insets are representative images of the top-down views of respective samples at higher magnification.

Figure 1. FE-SEM images of (a) TE-PET cross section revealed by FIB milling, (b) nanocone array prepared via Au electroless plating, and (c) TMA prepared with a gold-sputtered seed layer and 250 cycles of Ni PCD. Insets are representative top-down views of the respective samples at higher magnification.

seed layer into the template pore (Figure 2c). Therefore, the dimensions of these tall TMAs follow the statistical distribution of the template pore (see above). Characterization of Ni Deposits Seeded with Pt ALD. Parts a and b of Figure 3 show X-ray photoelectron spectroscopy (XPS) data of a flat PET membrane (i.e., without track-etched pores) treated with Pt ALD and Ni PCD, respectively. Figure 3a shows the Pt 4f XPS region after Pt ALD. The two peaks at 73.6 and 76.9 eV indicate the formation of Pt oxides, and the shoulders on the peaks suggest that multiple oxidation states are present.52 Pt oxide formation at deposition temperatures similar to our protocol have been attributed to the decomposition of the Pt precursor.51 No peaks for Pt metal were observed; however, Pt oxide thin films deposited by ALD have been shown to exhibit metallic conductivity.53 The Ni 2p XPS region after Ni PCD (Figure 3b) shows characteristic peaks for metallic Ni at 851.7 and 868.9 eV with little or no oxide present.54 This contrasts with the previous work by Göransson et al., who report the presence of NiO and Ni(OH)2 species in their Ni PCD sample.55 We attribute this discrepancy to the difference in the PCD

diameters (o.d.’s) of 1.07 ± 0.06 and 0.59 ± 0.07 μm, respectively. These dimensions are comparable to the TMAs prepared using only the gold-sputtered seed layer and Ni PCD (Figure 1c), and suggest that the penetration depth of the Pt ALD seed layer is similar to or smaller than that of the sputtered Au. On the other hand, when the Pt precursor was pulsed for 5 s (Figure 2b), the microtube heights were measured to be 3.94 ± 0.46 μm, with base and tip o.d.’s of 1.10 ± 0.05 and 0.46 ± 0.08 μm, respectively. These microtubes are indeed taller than those prepared by the sputtered Au seed layer (Figure 1c), yet shorter than the original template pore (Figure 1a). However, the base o.d. is still consistent with the original template pore, indicating that the Pt-ALD-seeded Ni deposit also shows high fidelity toward the original template geometry. The top-down view of a single microtube shown in the inset of Figure 2b demonstrates a smaller tip o.d. than that of shorter microtubes (Figures 1c and 3a). The hollow aspect will be discussed in detail below. When the precursor pulse duration was further increased to 10 s, the resulting TMAs exhibited sharp, closed tips, indicating full penetration of the Pt C


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Figure 3. (a) XPS of Pt ALD film on PET. (b) XPS of Pt ALD and 250 cycles of Ni PCD on PET. (c) EDX spectra of a TMA taken with top-down (upper trace) and bottom-up (lower trace) orientations. (d) XRD pattern of 20 000 cycles of Ni PCD on Cu foil, with the JCPDS cards for Ni and Cu shown below.

parameters: the work by Göransson et al. used longer pulse durations and lower current density then what we applied in our study. Energy-dispersive X-ray (EDX) spectroscopy (Figure 3c) was used to assess the composition of the TMAs prepared with Pt ALD (5 s pulse duration), Au sputtering, and 250 cycles of Ni PCD. When the microtubes are viewed from top-down, we expect the Pt layer to be exposed, with Au and Ni layers underneath (see Scheme 1). Indeed, the EDX spectrum of the microtubes viewed from top-down (upper trace) shows distinct peaks attributed to both Pt and Au. A strong Ni peak is also present, indicating that the X-ray signal is collected from a volume that extends beyond the Pt/Au layers and into the thicker underlying Ni layer. On the other hand, in the spectrum obtained from the bottom-up orientation of the TMA (lower trace) the Pt and Au peaks are largely suppressed and the Ni peak dominates. Again, this is expected since the Pt/Au layers are buried beneath the much thicker Ni layer. X-ray diffraction (XRD) of the Ni layer electrodeposited by PCD confirms the deposition of polycrystalline Ni (Figure 3d). This sample was prepared by 20 000 cycles of PCD on a Cu foil substrate to ensure sufficient thickness for the XRD experiment. A Cu foil substrate was used because the thick Ni PCD layer on PET resulted in delamination of the deposit from the PET. Interestingly, the deposited Ni film exhibits an increased

magnitude for the (200) peak, in contrast to the report by Nasirpouri et al., where the (111) peak showed a much higher intensity than that for the (200) peak.49 Since Ni has the same face-centered cubic structure and similar cell parameters as the underlying Cu substrate (0.3523 and 0.3615 nm for Ni and Cu, respectively), we attribute this to the epitaxial growth of Ni on the Cu substrate that has a preferred orientation along the (200) crystallographic plane (Supporting Information Figure S1).56 Effect of PCD Cycle Number on TMA Dimensions. The effects of PCD on the dimensions of the TMAs are summarized in Figure 4. The images of TMAs observed from the base side (Figure 4a) clearly demonstrate the decrease in pore opening with increased PCD cycles. From the corresponding measurements of the pore opening [i.e., the microtube base inner diameter (i.d.)] (Figure 4d), we found that the thickness of the deposit increases linearly with cycle number at a rate of 0.37 nm per cycle for the first 1000 cycles (coefficient of determination 0.961). Parts b and c of Figure 4 are side-view and top-down images taken from the tip side of the TMAs. The base o.d.’s measured from these images (Figure 4e) are independent of the PCD cycling, as expected, since they are defined by the dimensions of the template. The tip o.d.’s, shown in Figure 4f, are correlated to the heights (Figure 4g) by the conical shape of the template pores. The effect of PCD D


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Figure 4. FE-SEM images of TMAs taken from (a) beneath the arrays (i.e., bottom-up), (b) tip side at ∼90° angle, and (c) top-down. The columns correspond to 250, 500, 1000, 1500, and 2000 cycles of Ni PCD (left to right). Microtube dimensions with number of Ni PCD cycles: (d) base-side i.d., (e) base o.d., (f) tip o.d., and (g) height. All samples were prepared with Pt ALD using a 5 s Pt precursor pulse duration.

channels, and the slope of the current−voltage data was used as a measure of the lumen resistance. Figure 5d is the current− voltage data obtained for samples prepared with different Ni PCD cycles. An expanded view of the data at lower voltages is shown in the inset. Assuming Ohm’s law, the ionic resistances of the TMAs were calculated and plotted in Figure 5e (filled points). As expected, the ionic resistance increases with increasing PCD cycles, which is indicative of the correspondingly increasing wall thickness and decreasing lumen size. These data also indicate that the tip of the microtubes remains open even after 2000 cycles of Ni PCD, despite the constricted tip observed in the milled microtube at 500 cycles (Figure 5b). The tip i.d.’s can be calculated using the microtube dimensions (base i.d. and height) obtained from FE-SEM image analysis and the ionic resistance, R, using57

cycling on TMA height, and hence also tip o.d., was negligible; the microtubes remain shorter than the depth of the template pore even after 2000 cycles of Ni PCD. This shows that our fabrication technique provides control over TMA height via the Pt ALD step, and not via the template pore depth or the Ni PCD cycles. Effect of PCD Cycle Number on the Microtube Lumen. The inner structure of the TMA, partially exposed by FIB milling (Figure 5a), demonstrates the uniform deposition of Ni PCD along the pore wallcrucial for fabricating a hollow structure without clogs. The TMA in Figure 5a was fabricated with 5 s pulse duration of Pt ALD followed by Au sputtering and 250 cycles of Ni PCD. The dimensions of the milled microtube are consistent with what we report in Figure 4d: 164 nm for the wall thickness (accounting for the 54° tilt angle of the sample) and 128 and 820 nm for the tip and base i.d.’s, respectively. When the Ni PCD was increased to 500 cycles (Figure 5b), the wall thickness increased to 283 nm, and the lumen became visibly constricted at the tip end. The base i.d. here is 0.59 μm which is also similar to what we found in Figure 4d. The small discrepancy may be attributed to the possible deformation of the TMA during the FIB milling step. Ion transport through the TMA was studied to investigate the effect of Ni PCD on the microtubes’ tip i.d., which was too small to be measured from FE-SEM images. Briefly, the TMA was sandwiched between two half-cells filled with aqueous Cu2+ electrolyte, and Cu wires were immersed in each half-cell to act as electrodes (Figure 5c). A potentiostat was used to sweep the voltage, thereby driving ionic current through the microtube

d tip = L /(Rσπdbase)

(1)

where σ is the solution conductivity and dtip, dbase, and L are the tip i.d., base i.d., and height of the microtube, respectively. Using measurements of the microtube shown in Figure 5a and the ionic resistance of 77.6 Ω, σ is estimated to be 5.04 S m−1. Using this value in conjunction with eq 1, the heights from Figure 4g, and the base i.d.’s from Figure 4d, dtip under each PCD condition can be calculated (Figure 5e, open points). These data suggest that dtip decreases rapidly until ∼1000 cycles of PCD, after which its change becomes negligible. Together with decreased change in the base i.d. beyond the 1000th cycle discussed in Figure 4d, these observations indicate that the lumen size becomes relatively independent of the cycle number E


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Figure 5. FE-SEM images of FIB cross sections of microtubes after (a) 250 or (b) 500 cycles of Ni PCD. (c) Schematic of ionic-current measurement setup, (d) i−V measurements from TMAs prepared with different Ni PCD cycles, and (e) corresponding ohmic resistances and calculated tip i.d.’s. Inset of panel d is an expanded view at lower currents to show the difference in slope of the samples with thick Ni layers. All samples were prepared with Pt ALD using a 5 s Pt precursor pulse duration.

beyond the 1000th cycle. This may be attributed to limited Ni2+ transport into the lumen through the decreased base opening with longer deposition.27 Intracellular Delivery of Quantum Dots (QDs) into C. reinhardtii via TMAs. Recent studies have demonstrated that arrays of microtubes allow membrane-impermeable molecules and QDs to be injected into cells through the microtubes’ lumen.38−41 However, this method has never been developed for microalgal cells that are protected by a rigid cell wall barrier and are approximately the size of the nucleus of mammalian cells (∼10 μm). The TMA offers a unique opportunity for injection into microalgal cells due to its small dimensions: the larger bases of the microtubes give mechanical stability for penetrating through the cell wall without breakage, and the sharp tips allow for effective cell impalement.58 Indeed, we have previously demonstrated that a centrifugal force of 2000g enables the successful impalement of solid (i.e., not hollow) nanocones through the cell wall of the wild-type microalgae C. reinhardtii.33 A TMA was prepared with a 5 s Pt precursor duration for Pt ALD and 250 cycles of Ni PCD to demonstrate its potential application as a microinjection platform for the small microalgal cells. An aliquot of Tris−acetate−phosphate (TAP) medium containing ∼1 million C. reinhardtii cells was layered onto a 30% Percoll solution on the TMA, then the cells were impaled onto the TMA under centrifugation for 5 min at 2000g. A representative FE-SEM image of a microalga pierced by the

TMA is shown in Figure 6a. Note that the microtube is not broken nor bent upon impact with the hard cell wall of the wild-type C. reinhardtii. After the centrifugal deposition of microalgae onto the TMA, the array was placed over a reservoir containing 10 nm diameter CdZnS/ZnS core/shell QDs in deionized water (see Supporting Information for details on QD synthesis and Supporting Information Figure S2 for QD images and spectra). The QDs were allowed to diffuse through the TMA for 1 h, then the array was disassembled and lightly rinsed. After rinsing, the cells on the TMA were fixed with glutaraldehyde and mounted with Mowiol antifade mounting medium. The sample was imaged with confocal laser scanning microscopy (CLSM) using 405 nm laser excitation. This wavelength excites both the QDs and the fixed C. reinhardtii cells, as depicted by the photoluminescence spectra shown in Figure 6b. The QDs were specifically designed to exhibit a strong emission peak at 432 nm, in contrast to the cells that exhibit a strong emission peak at 670−680 nm with weaker emission peaks near 460 and 600 nm. By using QDs with a large difference in peak emission wavelength from the microalgal cell, location of the QDs can be identified without the need to filter false-positive signals from the microalgal autofluorescence molecules (e.g., chlorophylls and carotenes). Figure 6c shows representative single-plane CLSM images of the TMA-pierced C. reinhardtii after QD delivery. The left and center images are filtered for blue emission and far-red F


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Figure 6. (a) FE-SEM image of C. reinhardtii cells pierced on a TMA. (b) Photoluminescence (PL) spectra of QDs (blue trace) and C. reinhardtii (red trace), and single-plane confocal laser scanning microscopy (CLSM) images of (c) TMA-pierced microalgae after 1 h of QD delivery and (d) cells exposed to the same quantity of QDs without a TMA. The TMAs were prepared using a 5 s Pt precursor pulse duration for Pt ALD and 250 cycles of Ni PCD.

emission, respectively. The overlay of the two images (image on the right) clearly demonstrates the colocalization of the blue QD emission and the red cell emission, indicating successful internalization of the QDs into the cell. Corresponding z-stack images can be found in Supporting Information Figure S3. Careful observation of the left image also shows that the blue signal is not uniformly distributed throughout the cells, but is rather excluded from certain regions within the cells. This is indicative of the QDs being delivered into the cytoplasm while being excluded from intracellular subcompartments such as nuclei and pyrenoids.59,60 Such nonuniform distribution has also been confirmed from QDs delivered into human mesenchymal stem cells, and demonstrates the difficulty of QDs to bypass the membranes of subcellular compartments.25,61−63 The blue signal seen outside the cells can be attributed to QDs that diffused out from the open microtubes (i.e., not impaling cells) and aggregated or adsorbed onto adventitious contaminants in the solution. The efficiency of QD

delivery (number of cells with internalized QDs/number of cells observed) is estimated to be 9.8% from the CLSM images. To assess the possibility of spontaneous QD uptake by the cells without the TMA, CLSM images were also taken for a sample where QDs and cell suspension were mixed directly for 1 h (Figure 6d). The concentration, volume, fixation, and mounting procedures were identical to the samples prepared for Figure 6c. Here, the blue signal is only observed in the space between the algal cells, and no overlay of the blue QD signal with the red chlorophyll was observed. This confirms our hypothesis that the blue fluorescence seen within the cells in Figure 6c is due to QDs delivered through the TMA, and not due to direct uptake by the cells. It should be noted that the control sample exhibited strong blue emission due to clusters of aggregated QDs, and the images had to be collected using a lower photomultiplier tube voltage. Such significant clustering was not observed in the TMA sample in Figure 6c due to the lower quantity of QDs that diffused through the microtubes and into the cell suspension. Images of the control sample G


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Au Electroless Plating. Au was deposited into the TE-PET template pores as described previously.50 Briefly, the sample is soaked in methanol for 5 min, then immersed in a solution containing 0.5 g of SnCl2·2H2O, 0.5 mL of trifluoroacetic acid, 50 mL of methanol, and 50 mL of deionized water for 45 min. The sample is rinsed twice by immersion in methanol for 2.5 min each, then immersed for 7.5 min in a solution containing 0.6 g AgNO3 in 100 mL of water with NH4OH added dropwise to turn the solution first brown then clear. After two more 2.5 min soaks in methanol, the sample is immersed in the Au plating solution overnight, then rinsed with water and dried. The Au plating solution contains 0.07 g of NaHCO3, 0.53 g of Na2SO3, 1.83 mL of HCHO, 0.83 mL of Oromerse Part B replenishing solution (Technic, Inc.), and 33 mL of water, with the final pH adjusted to 10.0 with 1 M H2SO4. Prior to template removal and FE-SEM imaging, the Au layer on the tip-side surface of TE-PET was removed by gentle wiping with several cotton swabs soaked in isopropyl alcohol. Conductive Seed Layer Deposition on TE-PET. Before depositing the conductive seed layer, the tip-side surface of the TEPET was masked with a sacrificial layer of 100 nm Cu deposited by electron beam evaporation using a Lesker PVD 75 instrument. A quartz crystal microbalance was used to determine the Cu thickness. After the Cu masking step, ALD of 15 nm Pt was performed using a Cambridge NanoTech Savannah 100 instrument with the chamber set to 130 °C. Trimethyl(methylcyclopentadienyl)platinum(IV) precursor was obtained from Sigma-Aldrich and heated to 70 °C during deposition. Compressed oxygen was sent through a Del Ozone LG-7 ozone generator operating at 100% capacity, which provides ∼10% O3 according to the manufacturer’s specifications. After pumping down the ALD chamber, 5 sccm N2 was introduced as a carrier gas and the O3/O2 line was primed by pulsing it into the chamber several times. Successive ALD cycling was performed under a quasi-static mode,45,67 where each cycle was performed as follows: Pt precursor was pulsed for 2.5 s followed by another 2.5 s soak (5 s total Pt pulse duration) with the exhaust valve closed, then the chamber pressure was brought down to the base level by opening the exhaust valve for about 8 s, the N2 flow was stopped before the O3/O2 gas was pulsed for 5 s followed by another 3 s soak, and finally the chamber was pumped to its base level again and N2 flow was returned to 5 sccm. This cycle was repeated 333 times to deposit approximately 15 nm of Pt, based on a previous report.51 For deep penetration of the Pt layer, a 5 s pulse and 5 s soak (10 s total) were used instead of the 2.5 s pulse and 2.5 s soak described above. For shallow penetration of the Pt layer, a 0.5 s pulse and 0.5 s soak (1 s total) were used. Sputter deposition of Au was performed with a 200 W argon plasma using a Lesker PVD 75 with a radio frequency energy source. The deposition rate was determined beforehand to be 6.3 Å s−1, and 30 nm of Au was deposited. The sacrificial Cu layer and the Pt layer deposited on the tip-side surface of the TE-PET were removed by first wiping the surface with cotton swabs soaked in 1 M H2SO4, then soaking for 20 min in 1 M HNO3 with slow agitation on a rotator. The sample was rinsed with deionized water and stored dry until use. Ni PCD. A Princeton Applied Research VersaSTAT 4 potentiostat was used to deposit Ni onto the TE-PET coated with the Pt/Au seed layer. Ag/AgCl electrode (BASi) was used as the reference electrode, and Ni wire as the counter electrode. Watt’s Bath, containing 265 g L−1 NiSO4·6H2O, 48 g L−1 NiCl2·6H2O, and 31 g L−1 H3BO3, was heated to 45 °C and used as the electrolyte. The current was pulsed at −50 mA cm−2 for 1 ms followed by 0 mA cm−2 for 99 ms. The number of pulse/rest cycles (100 ms each) was varied between 250, 500, 1000, 1500, and 2000. Following the Ni PCD, all samples were thoroughly rinsed with water and dried under a stream of N2. TE-PET Complete Removal by Dissolution. Prior to TMA imaging with SEM, the samples were attached to Cu foil substrates using double-sided conductive Cu tape (3M). The TE-PET was then removed by immersion in 1,1,1,3,3,3-hexafluoroisopropyl alcohol (HFIP) (Oakwood Products, Inc.) for 1 h with slow agitation on a rotator. The samples were then rinsed gently with fresh HFIP and airdried. TE-PET Partial Removal by Reactive Ion Etching (RIE). A thin layer of Series 120 thermal joint compound (Wakefield Solutions) was

collected with identical conditions to that of Figure 6c are shown in Supporting Information Figure S4. Delivery of QDs into cells has been identified as a major limitation in using QDs as fluorescent probes for intracellular imaging,61 especially for algal species with cell walls where endocytosis is not a common mechanism for material internalization.64,65 Our results show that the application of a TMA prevents QD aggregation and enables a direct approach for delivering this fluorescent nanomaterial into the intracellular environment.

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CONCLUSION This study demonstrates the synergistic use of ALD and PCD to prepare template-synthesized TMAs with high aspect ratio and nanometer-sized tips. The TMA is prepared using conically shaped template pores with a wide base opening that narrows down to a sharp, closed tip. We demonstrate that, by decreasing the Pt precursor pulse duration from 10 to 1 s during the ALD step, the heights of the microtubes extending from the base can be controlled from the maximal full length (∼6 μm) to only a fraction (1−2 μm) of the template pore. The difference in the height is attributed to the limited penetration depth of the conductive Pt seed layer at shorter precursor pulse duration. Ni PCD results in smooth deposition of Ni within the template pores with a thickness that can be controlled as a function of PCD cycle. Ionic current measurements confirm that the microtubes’ lumen stayed open even after 2000 cycles of Ni PCD. Potential application of this platform as a microinjection device is demonstrated using C. reinhardtii with intact cell walls. Cells that are impaled by the TMA exhibit distinct fluorescence from membrane-impermeable CdZnS/ZnS QDs, indicating successful intracellular delivery of the QDs through the TMA. QDs that are added directly to the external cell medium experience significant aggregation and are not internalized by the cells. The TMA platform described in this study not only enables QD-based imaging techniques to be more accessible for microalgal studies, but also offers opportunities for studying nanoparticle-augmented light harvesting, studies on nanomaterials’ intracellular toxicity, and biosensing with microalgae. Although beyond the scope of this work, this platform may also be developed for imaging molecular functions of live microalgal cells, based on prior studies that report cell survival after impalement with microtube arrays.38−41,66

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METHODS

Materials. Unless otherwise noted, all chemicals were purchased from Fisher Scientific and used as received. Deionized water was obtained using a Milli-Q Integral water purifying system (18.2 MΩ cm). Template Track Etching. Ion-tracked PET membranes were obtained from GSI, Darmstadt, Germany. Irradiation was done with Au25+ ions at 10.4 MeV with 106 cm−2 fluence. The track-etch procedure was done as described previously.31,33 Briefly, the membranes were first irradiated with 254 nm UV light for 1 h per side. Then the membrane was placed between two half-cells, one halfcell containing 1 M HCOOH and 1 M KCl in water and the other containing 4 M KOH in methanol. A Pt wire electrode was placed in each half-cell and a positive bias of 60 V was applied to the electrode in the KOH solution during the etching process. The etch proceeded for 250 s, after which the KOH solution was quickly replaced with the HCOOH solution for 10 min. After rinsing with water, the etched membrane was soaked in water for several hours, then dried. H


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ACS Applied Materials & Interfaces used to hold the sample on the cooled (23 °C) electrode in the RIE chamber. The etching conditions were as follows: 30 sccm O2, 100 W radio frequency power with minimal (0−2 W) reverse power, 100 Pa chamber pressure (obtained by partially closing a throttle valve on the vacuum line), and a total etch time of 24 min. Thermal joint compound was removed by thorough rinsing with hexane and acetone, followed by successive immersions in water (several times), 0.1 M H2SO4 with slow agitation on a rotator (5 min), water (5 min), and isopropyl alcohol (1 min). Finally, the sample was dried and stored until use. Characterization. XPS was done using a Kratos Axis Ultra XPS system with a monochromated Al Kα source (1486.6 eV). The spectra were analyzed with CasaXPS software and calibrated to the adventitious C 1s signal (285.0 eV). XRD was done using PANalytical X’Pert multipurpose diffractometer using Cu Kα radiation (λ = 1.5406 Å). FE-SEM, FIB milling, and EDX spectroscopy were done using a Zeiss Auriga dual-beam FIB-SEM. Microtube dimensions were obtained from FE-SEM images using ImageJ software.68 Ionic currents were measured using a Princeton Applied Research VersaSTAT 4 potentiostat and an aqueous electrolyte containing 0.8 M CuSO4, 0.92 M H2SO4, and 3.33 mM HCl. Confocal laser scanning microscopy was performed using an Olympus FV1000 inverted microscope equipped with a 405 nm laser line. A 60× oil immersion lens and a zoom of 5× were used for imaging. QD Synthesis. CdZnS/ZnS core/shell QDs were synthesized by modifying an established procedure.69 They were capped with glutathione for aqueous transfer based on a method reported previously.70 Details of the synthesis, transmission electron micrographs, and photoluminescence data are included in the Supporting Information. Microalgae Cultivation. The wild-type microalgal species C. reinhardtii (catalog number CC124) was obtained from the Chlamydomonas Resource Center at the University of Minnesota. The cells were grown in TAP medium with constant agitation and an alternating 12 h light/12 h dark cycle. The cells were collected during log-phase growth. Cells were counted using a Bio-Rad TC10 automated cell counter. Centrifugation of Microalgae onto TMAs. The TMA samples were first degassed in deionized water under rough vacuum for 30 min, then transferred to a custom 3-D printed acrylonitrile−butadiene− styrene device. An amount of 300 μL of 30 vol % Percoll (GE Healthcare) in TAP medium was pipetted onto the TMA, followed by another layer of TAP medium containing 106 microalgal cells (usually 30−80 μL). After the device was centrifuged at 2000g for 5 min, the supernatant was carefully removed with a micropipette, the device was disassembled, and the TMA was transferred to a different custom device for QD delivery. The delivery device consisted of a poly(dimethylsiloxane) well that was bonded to a glass microslide via UV-ozone treatment (PSD-UV instrument, Novascan Technologies, Inc.) and baked at 70 °C for 1 h. The chamber contained 75 μL of 4.8 μM QDs in water, onto which the TMA was carefully placed with the base side facing the QD solution. An amount of 750 μL of TAP was added on top of the TMA with the centrifuged cells, and the QDs were allowed to diffuse through the TMA for 1 h. Finally, the delivery device was disassembled, the TMA was gently rinsed, and the cells were fixed following the procedure outlined below. As a control, the same quantity and concentration of microalgae cells and QDs were mixed on a glass coverslip and allowed to sit for 1 h before fixing. Fixation of Microalgae. The TMA with centrifuged cells was first rinsed twice with 250 μL of 80 mM cacodylate buffer pH 7.4 for 5 min each, then treated with 2 vol % glutaraldehyde in cacodylate buffer for 1 h at 4 °C. Following two more rinsing steps with cacodylate buffer, the TMA was mounted on a glass coverslip with Mowiol antifade mounting medium (24 g of glycerol, 9.6 g of Mowiol 4-88 (SigmaAldrich), 48 mL of 0.2 M Tris buffer pH 8.5, and 24 mL of deionized water). A coverslip was placed above the medium and sealed with clear nail polish to prevent drying. Samples were stored at 4 °C in the dark prior to imaging with CLSM.

Sample Preparation for FE-SEM Imaging. After the final rinsing of the fixed cells described above, the samples were rinsed with deionized water for 5 min, then dehydrated with 10%, 30%, 50%, 70%, 90%, and 100% ethanol solutions for 10 min each. A Samdri-PVT-3B critical point dryer (Tousimis) was used to exchange the ethanol with CO2. Samples were sputter-coated with ∼10 nm Au and kept in a desiccator prior to imaging with FE-SEM.

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ASSOCIATED CONTENT

S Supporting Information *

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsami.6b11062. XRD pattern of Cu substrate, QD synthesis details, transmission electron micrographs and spectral analyses of QDs, and CLSM images of control sample imaged with high photomultiplier tube voltage (PDF)

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. ORCID

Todd D. Krauss: 0000-0002-4860-874X Hitomi Mukaibo: 0000-0001-7195-6403 Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS The research has been conducted under the financial support from the University of Rochester, NSF DGE-0966089, and NSF CHE-1307254. We are grateful to B. McIntyre and the Integrated Nanosystems Center for use of FIB-SEM, XPS, PVD, and ALD instruments, to C. Pratt for XRD measurements, to L. Callahan and the Light Microscopy Core for CLSM, to J. McGrath for 3D printing, and to J. Miller and the River Campus Instrument Machine Shop for consultation and fabrication of the U-cell device.

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REFERENCES

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