Preparation of Mica and Silicon Substrates for DNA Origami Analysis and Experimentation

Summary

Reproducible cleaning processes for substrates used in DNA origami research are described, including bench-top RCA cleaning and derivatization of silicon oxide. Protocols for surface preparation, DNA origami deposition, drying parameters, and simple experimental set-ups are illustrated.

Abstract

The designed nature and controlled, one-pot synthesis of DNA origami provides exciting opportunities in many fields, particularly nanoelectronics. Many of these applications require interaction with and adhesion of DNA nanostructures to a substrate. Due to its atomically flat and easily cleaned nature, mica has been the substrate of choice for DNA origami experiments. However, the practical applications of mica are relatively limited compared to those of semiconductor substrates. For this reason, a straightforward, stable, and repeatable process for DNA origami adhesion on derivatized silicon oxide is presented here. To promote the adhesion of DNA nanostructures to silicon oxide surface, a self-assembled monolayer of 3-aminopropyltriethoxysilane (APTES) is deposited from an aqueous solution that is compatible with many photoresists. The substrate must be cleaned of all organic and metal contaminants using Radio Corporation of America (RCA) cleaning processes and the native oxide layer must be etched to ensure a flat, functionalizable surface. Cleanrooms are equipped with facilities for silicon cleaning, however many components of DNA origami buffers and solutions are often not allowed in them due to contamination concerns. This manuscript describes the set-up and protocol for in-lab, small-scale silicon cleaning for researchers who do not have access to a cleanroom or would like to incorporate processes that could cause contamination of a cleanroom CMOS clean bench. Additionally, variables for regulating coverage are discussed and how to recognize and avoid common sample preparation problems is described.

Introduction

First introduced in 2006, DNA origami utilizes the self-assembling nature of DNA oligonucleotides to produce designable and highly ordered nanostructures.¹ A myriad of structures have been reported, ranging from smiley faces to latched 3-dimensional boxes.² DNA origami can be functionalized with various biomolecules and nanostructures, giving rise to research applications in nanoelectronics, medicine, and quantum computing.³ However, the analysis and many future applications are not only dependent on structural design, but also on the adhesion of the DNA origami nanostructures to surfaces. The methods described in this manuscript pertain to the preparation of DNA origami samples on two types of substrates: mica and functionalized silicon oxide.

Mica is the substrate of choice for DNA origami studies because it is atomically flat, with a layer height of 0.37 nm ± 0.02 nm.⁴ It is also easily cleaned, making sample preparation and atomic force microscopy (AFM) studies straightforward. Muscovite mica contains a high density of potassium in each cleavage plane, but these ions diffuse away from the mica surface when in water. To mediate the binding of DNA origami to the mica substrate, Mg²⁺ is used to reverse the negative charge of the mica and electrostatically bind the DNA phosphate backbone to the substrate (Figure 1A).⁵ Mixtures of annealed DNA in the presence of large excesses of staple strands give high coverage and good images on mica because the adhesion of DNA origami to the Mg²⁺-terminated surface is much stronger than the adhesion of single-stranded oligonucleotides (staple strands). Other positively charged ions, including Ni²⁺ and Co²⁺ can be used to control the adhesion of DNA on mica.^6,7 Changing the concentration of monovalent and divalent cations in solution can mediate adhesion and surface diffusion rates of DNA origami.⁸ However, the protocol for preparing mica substrates and depositing and rinsing the origami is often not explicitly described in published manuscripts.⁹ Without a clear protocol, reproducible results can be difficult to obtain.

Mica is an insulator, so it is not suitable as a substrate for some applications in nanoelectronics. Silicon passivated with a thin native oxide has desirable electronic properties, including compatibility with prior complimentary metal-oxide semiconductor (CMOS) processing to create input/output structures and topographic features. Silicon wafers stored in air are passivated with either a thick thermal oxide or thin native oxide film that is relatively dirty, with a high particulate count. Silicon oxide has a much lower surface charge density than mica, and the charge density is highly dependent on oxide preparation and history. At magnesium ion concentrations above 150 mM, good coverages (up to 4/µm²) of rectangular DNA origami can be achieved on oxygen plasma treated silicon substrates; however, this concentration and coverage may change depending on the size and design of the nanostructures being used.¹⁰ An alternative protocol for tuning the surface charge is to attach a cationic self-assembled monolayer of 3-aminopropyltriethoxysilane (APTES) (Figure 1B) to the oxide. The primary amine on APTES can be protonated at pH values below 9, modifying the charge and hydrophobicity of the substrate.¹¹ For a complete monolayer of APTES to be successfully deposited, the silicon must be appropriately cleaned using Radio Corporation of America (RCA) protocols. These protocols include treatments in ammonium hydroxide and hydrogen peroxide solutions (RCA1) to remove organic residues and particle contaminants. A short etch in aqueous hydrofluoric acid solution removes the native oxide layer along with any ionic contaminants that adhere to the oxide. Finally, samples are exposed to a hydrochloric acid and hydrogen peroxide solution (RCA2) to remove metal and ionic contaminants and form a thin, uniform oxide layer.¹² Most cleanrooms have designated hoods for CMOS cleaning protocols, with strict rules about what can be used in these areas. A common problem comes in the form of ions such as sodium, which can disrupt the electronic properties of CMOS structures by creating midbandgap traps.¹³ Ions commonly used in DNA origami preparation and deposition buffers could contaminate the CMOS baths and cause problems for other researchers using the clean room. For this reason, our group uses a 'dirty' CMOS cleaning bench arranged specifically for the small samples used for DNA origami research. This process is a good alternative to the traditional cleanroom set-up and may be suitable for laboratories that do not have access to a cleanroom CMOS bench.

Protocol

1. Experiment Planning and Material Preparation

1. Determine the design, concentration, and functionality of the DNA origami that will be used in the experiments.^14-16 Here, we use a DNA origami rectangle design prepared in 1x TAE/Mg²⁺ solution (40 mM Tris-base, 20 mM acetic acid, 2 mM EDTA and 12 mM magnesium acetate, pH 8.0).¹⁷
2. Autoclave all tips, tubes, and containers to be used. These materials must all be autoclave compatible.
3. Prepare a supply of sterile water for rinsing. Fill a sterile jar with approximately 500 ml of 18 MΩ x cm water, place on a hotplate, boil for 5 min with the cap off, and take the jar off of the hotplate and let cool with the lid placed on the container but not tightened. Store in the refrigerator and prepare a new supply each month or when necessary.

2. Preparing the Mica Substrate

1. Cut substrates to the appropriate size (1 cm x 1 cm squares) with scissors. Mica is thin and brittle. Alternatively, purchase mica in disc form that does not require cutting.
2. Cleave the mica using double-sided tape. Mica is composed of layers of minerals separated by intercalating ions, each layer can be peeled off when adhered to double-sided tape.¹⁸
   1. Place the mica squares on the double-sided tape still in the tape dispenser, making sure it is adhered firmly to the tape. Carefully slide the tweezers between the mica and the tape, the top-most layer will be removed and remain on the tape. Immediately after the removal, the mica square will have its clean side facing downward. Make sure to flip the mica over before storing in a container.
   2. Repeat this three to four times to ensure complete removal of the top most layer and adequate cleaning.
3. Alternatively, adhere the mica to a container or table-top with a piece of double sided tape and use a second piece to stick to and peel off the top most mica layer. The mica will be properly cleaned in both cases, although the alternative method makes depositing DNA and rinsing and drying the sample difficult due to the second adhesion to the support.

3. Depositing DNA Origami on Mica

1. Briefly, mix the vial of DNA origami using a vortex mixer to ensure even dispersal of nanostructures in solution.
2. Pipette 4 µl of solution onto the mica, ensuring that the pipette tip does not touch the substrate. Leave the DNA origami on the mica for approximately 10 min to ensure adequate coverage. The deposition time will vary depending on the concentration of DNA origami used as well as the desired coverage (Figures 2 and 3).
   1. Rinse the DNA origami solution off of the mica substrate using 100 µl of sterile water over a sink or other liquid receptacle. Pick up the mica using tweezers. Pipette the water on the substrate, with the flow of the drop towards the tip of the tweezers. Shake the mica with a sharp motion downwards to remove the excess water. Hold the tweezers upright so that the water will flow toward the tweezers to avoid contamination of the sample.
3. Dry the substrate with a steady stream of nitrogen (N₂) for 1 min. Make sure that any excess water is removed. Repeat the rinse with an additional 100 µl of sterile water. Dry the substrate with N₂ for an additional 3 min. A completely dry substrate is necessary for successful atomic force microscopy (AFM) analysis (Figure 4).
4. Analyze the substrate using AFM or store in a closed container.

4. CMOS/Silicon Cleaning Set-up

1. CAUTION: When using the CMOS set-up, use personal protective equipment at all times. The reagents include strong acids, strong bases, hydrofluoric acid (HF), and strong oxidizing agents which can react with waste solvents if reagents are not properly disposed of. Adhere to the following safety precautions:
   1. House the CMOS bench in a chemical hood with no other processes or set-ups.
   2. Wear nitrile gloves, lab coat, safety goggles, large industrial nitrile gloves, a spill apron, and a face shield at all times when using the CMOS bench.
   3. Use plastic tubs as secondary containment when solutions are prepared.
   4. Use an inert fluorinated polymer measuring beaker for handling the concentrated HF.
   5. Make calcium gluconate ointment available as first aid for any skin exposure.
   6. Only allow properly trained personnel to perform the process.
   7. Always ensure another member of the lab is present in case of emergency.
   8. Keep MSDS information for all chemicals near the hood.
   9. Be familiar with the institution's or company's chemical spill and exposure policies.
      Note: HF readily permeates skin and is a calcium scavenger, affecting bones and damaging nerves if exposure occurs. Dermal exposure to a few milliliters of concentrated hydrofluoric acid can be dangerous and even fatal. Establish necessary precautions to ensure exposure does not occur.
2. Perform RCA1 and RCA2 in a separate 250 ml glass beakers on separate hotplates. Each beaker should contain a stir bar. Monitor the temperature of the solution using thermometers clamped so the stir bar does not bang into the bulb. Cover the beakers using a watch glass to diminish the effects of evaporation.
3. RCA1 Preparation
   1. Place 50 ml of 18 MΩ x cm water into the designated RCA1 beaker using a measuring beaker.
   2. Add 15 ml of concentrated ammonium hydroxide (NH₄OH) to the beaker. Rinse the measuring beaker with 25 ml of water and add the rinse water to the RCA1 beaker.
   3. Turn on the heat and stirrer on the hotplate and bring the RCA1 bath to 70 °C.
   4. Add 15 ml of 30% hydrogen peroxide (H₂O₂) to the RCA1 beaker. Use the RCA1 solution within 1 hr after the H₂O₂ has been added. The bath can be used several times within the span of three days if 15 ml of peroxide is added to the bath each time.
   5. Rinse the measuring beaker thoroughly with water and discard the rinse in an appropriate RCA1 waste bottle.
4. RCA2 Preparation
   1. Add 70 ml of 18 MΩ x cm water to the designated RCA2 beaker using the thoroughly rinsed measuring beaker.
   2. Add 15 ml of concentrated hydrochloric acid (HCl). Rinse the measuring beaker with 20 ml of water and add it to the RCA2 beaker.
   3. Increase the heat and stir speed of the hotplate until the solution reaches 70 °C.
   4. Add 15 ml of 30% H₂O₂. Like the RCA1 bath, use this solution within 1 hr from when the H₂O₂ is added; additionally, the bath can be reused several times within the span of three days if 15 ml of H₂O₂ is added before each use.
5. HF Solution Preparation
   1. Place 50 ml of water in an inert fluorinated polymer beaker.
   2. Measure 4 ml of concentrated hydrofluoric acid (49%) in the plastic measuring beaker and add it to the inert fluorinated polymer beaker.
   3. Rinse out the plastic measuring beaker with a total of 50 ml of water, adding the rinse water to the HF beaker. Wash out the measuring beaker thoroughly with water and discard the washings in a designated HF waste container.

5. Preparing and Cleaning the Silicon Substrate

1. Cutting silicon wafers into chips
   1. Identify the perpendicular and parallel lattice directions on the flat polished surface of the silicon wafer. These directions are used to help make cleaving squares easier. The following instructions pertain to cleaving silicon <110> and may not be suitable for other crystal orientations.
   2. Place the silicon wafer polished-side-up on a soft surface, such as a napkin. Using the diamond-tipped scribe pen, gently nick the bottom of the wafer along the primary flat edge. Place a small wire, such as a paperclip, below the nick and gently apply pressure to the wafer by placing fingers or tweezers on either side of the nick and pushing down. Doing this will separate the wafer into two halves along the crystal lattice line in the natural cleave direction.
   3. On another napkin, with a pencil and ruler, measure out the desired width of the squares by marking dots on both the top and the bottom of the napkin. Connect these dots with straight lines. This will serve as a guideline for even square shapes.
   4. Place one of the wafer halves edge-first between the measured lines on the napkin flushed against the line and repeat the in step 5.1.2 The freshly broken perpendicular pieces should now be the width of the cleaved squares. Turn the wafer horizontally on the napkin. Place it between the perpendicular lines and repeat the process in step 5.1.2.
   5. Store the freshly cleaved wafer chips in a clean vial filled with DI water to prevent scratching. The silicon chips can be stored indefinitely, but should be cleaned before starting experiments.
2. CMOS Cleaning of Silicon
   1. When the RCA1 solution has reached the appropriate temperature, submerge eight to ten 1 cm x 1 cm silicon chips in the solution using an inert fluorinated polymer basket with a 2" diameter. Bubbles of oxygen will form on the chips and beaker walls. If no bubbling occurs, the H₂O₂ is degraded. Leave the chips in the solution for 10 to 20 min, agitating the basket up and down every few minutes to keep the chips from sticking together.
   2. Lift the basket containing the silicon chips up and drain well. Move the basket over to the waste beaker and rinse thoroughly with 18 MΩ x cm water. Immerse in the wash beaker and jiggle up and down for 20 sec. Drain the basket and rinse thoroughly with water over the waste beaker. Empty the waste beaker into a designated RCA1 waste bottle and refill with water.
   3. After RCA1 cleaning is complete, place the basket into the 1:50 HF beaker for 10 to 20 sec. Use a gentle up and down motion to mix the chips and HF. Lift the dunk bucket to allow the HF to drain completely away.
      Note: The chip surfaces should be hydrophobic; water will not wet the chip, but instead form droplets with high contact angles. This indicates that the silicon oxide has been etched away and the chip is now terminated by Si-H bonds.
   4. Place the wash beaker and rinse beaker in a plastic tub, move the basket over the rinse beaker and rinse with 18 MΩ x cm water. Submerge the basket into the wash beaker and agitate for 20 sec.
   5. Complete a second drain and rinse cycle with 18 MΩ x cm water. Dump the wash water into the rinse beaker and refill the wash beaker with water. Pour all waste into a designated plastic HF waste bottle.
   6. When the RCA2 solution has reached the appropriate temperature, submerge the silicon chips in the solutions using the basket. Leave in the solution for 10 to 20 min. The chip will now be hydrophilic due to growth of a thin (1-2 nm) oxide film.
   7. Remove after the appropriate amount of time and follow the same rinsing procedures as for RCA1. Dispose of the waste in the appropriate RCA2 waste container. Remove each chip from the basket with plastic tweezers, rinse with water, and blow dry with nitrogen.
   8. Store chips in a plastic wafer box or in a vial of 18 MΩ x cm water. The silicon will remain clean when stored in water for approximately three days after cleaning. Make sure that the work area is properly cleaned up and the exterior of the CMOS gloves have been washed. Leave the CMOS gloves in the hood to dry.

6. Depositing DNA Origami on APTES-functionalized Silicon

1. Self-assembled Monolayer Formation on Silicon
   1. Warm APTES to RT before opening. If the bottle is too cold, condensation may occur, causing hydrolysis of the APTES during storage. Add 1,980 µl of 18 MΩ x cm water and 20 µl of APTES to a clean scintillation vial and swirl to mix. Use this solution immediately.
   2. Place a cleaned silicon chip reflective-side-up in the scintillation vial, cap it, and let sit for 20 min. Remove the chip using tweezers and rinse with 200 µl of water and dry for 1 min with a stream of N₂.
2. Depositing DNA origami on APTES Functionalized Silicon
   Note: The steps for depositing DNA origami on functionalized silicon are analogous to those for depositing on mica.
   1. Briefly mix the DNA origami vial and pipette 4 µl of solution onto the silicon substrate. If necessary, increase the volume of DNA origami solution used to cover the entire substrate as the functionalized silicon is more hydrophobic than the mica substrate. Use a glass cover slip to press the deposition solution down and prevent evaporation during long depositions.
   2. Let the solution stand for the amount of time necessary for the concentration used and the coverage desired (see Figures 2 and 3 for the effect of time and concentration on surface coverage). Rinse the substrate with 100 µl of sterile 18 MΩ x cm water and dry with N₂ for 1 min.
   3. Repeat the rinsing with an additional 100 µl of sterile water and dry the substrate with N₂ for 3 min.
   4. Store the sample in a clean container until further experiments or imaging can be performed. Samples begin to show particulate accumulation after approximately one to two weeks of storage depending on how much they are handled.

7. AFM Imaging and Image Analysis of DNA Origami Samples

1. Use AFM in Tapping mode in air for imaging purposes. Tapping mode ensures that minimal force will be applied to the fragile nanostructures, compared to contact mode.
   Note: The imaging parameters will be dependent on the instrument. All images presented were captured using a MultiMode Nanoscope IIIa.
2. Select AFM probes for non-contact/tapping mode in air, with gold reflective coating, a resonant frequency (nominal) of ~300 kHz, force constant of 40 N/m, and tip radius <10 nm.
3. Process and analyze the AFM images using NanoScope Analysis Software. Perform coverage calculations using ImageJ.¹⁹

Representative Results

Two variables dictate the coverage of DNA origami on the substrate: solution concentration and exposure time. The adsorption characteristics of DNA origami on mica and APTES functionalized silicon oxide have been previously reported.¹³ The relationship between the concentration of DNA origami in the deposition solution and the final coverages on mica are summarized in Table 1 and Figure 2, showing increasing concentration results in increased coverage. The time-dependence of binding is seen in Figure 3. Surface coverage was previously studied to quantify the binding behavior of DNA origami on mica and modified silicon oxide surfaces. DNA origami in 12 mM magnesium in 1x TAE buffer has 83.3% ± 3.1% coverage on mica after 30 min absorption time on the surface. Maximum coverage on silicon oxide modified with APTES SAMs is observed after 60 min, which is less than the maximum coverage on mica. A longer deposition time is needed if a high surface coverage is required on APTES functionalized silicon oxide.

There are several variables that can cause poor sample preparation. The most troublesome is inadequate rinsing and drying. If the buffer solution is not rinsed properly, large aggregates form on the substrate (Figure 4A). 'DNA origami islands' are observed when the nanostructures adhere to patches of magnesium salts on the surface (Figure 4B). Finally, with high coverage samples, it is possible to have excess buffer components bridging between individual DNA origami (Figure 4C), which makes it difficult to differentiate nanostructures using AFM. These results can be avoided by following a thorough rinsing and drying protocol for both mica and silicon substrates.

Formation of APTES films on silicon oxide substrates can pose problems as well. Silicon wafers have a rough silicon oxide layer that must be removed and a smoother, thinner silicon oxide layer reformed before it can be functionalized. A properly cleaned silicon substrate is illustrated in Figure 5A. During cleaning of the silicon wafer, it is important to make sure the number of silicon chips is not too high, as it is possible for two chips to become stuck to one another, blocking exposure to the reagents (Figure 5B). If cleaned silicon has been stored in 18 MΩ x cm water for more than one week, the contaminant layer will reform and recleaning is necessary. The APTES supply may also cause problems with sample preparation. APTES readily polymerizes through hydrolysis, which is the basis for the monolayer formation.²⁰ The extent of this polymerization is dependent on the concentration of water that the APTES is exposed to. Over time and through repeated use, it is possible for water to condense inside the APTES bottle and contaminate the supply. The resulting polymerization produces large aggregates that adhere to the substrate (Figure 5C). The increased roughness and presence of aggregates makes identifying DNA nanostructures using AFM difficult. It is good practice to store the APTES bottle in a plastic bag in the refrigerator, and let the APTES bottle warm to RT before opening to avoid condensation.

Figure 1. A schematic comparing the binding mechanism for DNA origami on (A) mica and (B) APTES functionalized silicon oxide (not to scale). Binding with mica is mediated by the presence of divalent cations, usually Mg²⁺. A monolayer of the protonated amine terminated 3-aminopropyltriethoxysilane is used to promote adhesion on the silicon oxide substrate.

Figure 2. AFM images illustrating variable coverage after 10 min depositions of (A) 2 nM, (B) 4 nM, and (C) 6 nM on mica. The height scale for all images is 5 nm.

Figure 3. Trends in surface coverage for 2 nM DNA origami in 1x TAE buffer, 12 mM Mg²⁺. MICA = purple line and circle markers, APTES = yellow lines and triangle markers. N=3 in determination of standard error.

Figure 4. Poor rinsing and drying can cause (A) solution aggregation on the substrate, (B) DNA origami islands, and (C) bridging of nanostructures on high coverage samples in the presence of excess buffer salts. The height scale for all images is 5 nm.

Figure 5. (A) Clean silicon should have RMS roughness less than 0.5 Å over 1 square micron. A complete APTES SAM deposited on a good native oxide should have an RMS roughness less than 1 Å over 1 square micron. (B) APTES film formed on an incompletely cleaned silicon oxide with RMS roughness of 2.29 nm over 1 square micron. Note the gaps and roughness of the APTES film. (C) APTES surface formed from a sample of APTES contaminated with condensed waste; hydrolysis in the APTES bottle forms large particulates. The height scale for all images is 5 nm.

Table 1. Percent coverage measurements for mica substrates with varying DNA origami solution concentrations. All deposition times are 10 min.

DNA Origami Solution	% Coverage on Mica Substrate
2 nM	8.49 ± 2.67 (N=5)
4 nM	55.89 ± 5.65 (N=3)
6 nM	77.44 ± 1.89 (N=4)

Discussion

There are several steps that need to be emphasized to attain consistent and ideal results. For mica samples, following a strict and thorough rinsing and drying regime, as in steps 3.3 and 3.4, will ensure that high quality images of individual DNA origami can be attained using AFM without the various problems outlined in the Representative Results section. Of primary importance for silicon samples is the cleanliness of the substrate. Following the cleaning procedures outlined in step 5.2 thoroughly and meticulously will assure that an appropriately cleaned silicon oxide surface will be attained. Additionally, monitoring the quality and effectiveness of chemicals, such as the hydrogen peroxide, hydrofluoric acid, and APTES, will ensure that the procedure runs smoothly.

The techniques described are not limited to only aqueous solutions of APTES. A mixed monolayer of APTES and trimethylaminopropyltrimethoxysilyl chloride (TMAC) can be used to tune the silicon surface charge and promote variable DNA origami adhesion.¹¹ The TMAC contains a permanently charged terminal quaternary amine –N(CH₃)₃⁺, compared to the pH dependent charge on APTES. Because the solution environment does not affect the protonation state or the charge of TMAC, varying the solution concentration of TMAC can tune the surface charge of the mixed monolayer and affect the interaction between the substrate and the DNA origami. Optimal DNA origami binding was observed for monolayers having a surface charge of 0.75-1.5 charges/nm², which corresponds to SAMs containing 100% to 40% TMAC concentration. This optimal surface charge prompted DNA origami coverages of approximately 110 origami/µm² on APTES SAMs and 120 origami/µm² on TMAC SAMs.

One advantage of silicon is its compatibility with lithographic patterning processes. High magnesium concentrations can be used with plasma-treated silicon oxides to promote selective binding of DNA origami to silicon. Care must be taken when removing the sample from the deposition solution to avoid rinsing the magnesium ions away and deforming the DNA origami.^21,22 The APTES process forms covalently attached cations on the silicon oxide surface, so washing or rinsing with buffer or water does not damage the attached DNA origami. The 'molecular liftoff' method is another possible route for patterning silicon substrates and promoting DNA origami adhesion. The silicon substrate is patterned using electron beam lithography and APTES is deposited on the exposed substrate. Following liftoff of the photoresist, DNA origami can be deposited on the patterned surface, preferentially binding to the APTES.²³

The interaction of DNA origami with a substrate changes its stability, opening new avenues for research and applications. DNA origami adhered to mica can be heated to 150 °C without visible alteration of nanostructure dimensions and minimal chemical changes.²⁴ This is in stark contrast to the fragility of DNA origami in solution, where the nanostructures are complete dehybridized above 70 °C.^25,26 This stability is maintained on heated silicon oxide substrates.²⁷ Even in diverse solvent systems, such as hexane, toluene, and ethanol, the shape and coverage of the nanostructures are maintained. The surprising stability of DNA origami indicates that applications that were previously thought incompatible, such as plasma enhanced vapor deposition, use of common photoresists and solvents, and unique chemical deposition environments, may be used in conjunction with DNA origami. However, whether the DNA origami maintain their functionality is still unknown and may limit possible applications.

Although the stability of DNA origami at elevated temperatures and in a limited number of solvent systems has been determined, the long-term stability of DNA origami on substrates is still unknown. The use of sterile techniques is necessary to avoid possible contamination, but this cannot be avoided after surface deposition. Imaging and sample analysis must be completed almost immediately after sample preparation; if the sample is stored for too long (greater than a week) various examples of sample degradation are often identified, including particulate accumulation and broken DNA nanostructures. Possible research avenues in nanoelectronics, biosensing, and other substrate based DNA origami applications may be limited by time-dependent destabilization of the DNA. Identifying the limitations of these techniques for applications requiring stability for longer periods of time requires further investigation.

Materials

Name	Company	Catalog Number	Comments
Eppendorf epT.I.P.S. Reloads, capacity 2-200 μl	VWR International, LLC	22491733	10 reload tray of 96 tips
Microcentrifuge Tubes, Polypropylene	VWR International, LLC	87003-290	0.65 ml, natural
Research Plus Pippete - Single Channel - 20-200 μl	A. Daigger & Company, Inc.	EF8960F-3120000054 EACH	Adjustable Volume
Research Plus Pippete - Single Channel - 2-20 μl	A. Daigger & Company, Inc.	EF8960D-3120000038 EACH	Adjustable Volume
Scotch 237 Permanent Double-Sided Tape	Office Depot, Inc.	602710	3/4" x 300", Pack of 2
Vortex Mixer	Thermo Scientific	M37610-33Q
Wafer container single, 2" (50 mm), 60 mm x 11 mm	Electron Microscopy Sciences	64917-2	6 per pack
6" Wafer, P-type, <100> orientation, w/ primary flat	Nova Electronic Materials, Ltd.	GC49266
Powder-Free Nitrile Examination Gloves	VWR International, LLC	82062-428	Catalog number is for size large
High Accuracy Noncontact probes with Au reflective coating	K-Tek Nanotechnology, Inc.	HA_NC/15
Autoclave Pan	A. Daigger & Company, Inc.	NAL692-5000 EF25341C
Sol-Vex II Aggressive Gloves, Size: 9-9.5; 15 mil, 13 inch - 1 dz	Spectrum Chemical Mfg. Corp.	106-15055	Before use, rinse with water and scrub together until no bubbles form on the gloves.
Tweezers PTFE 200 mm Square	Dynalon Corp.	316504-0002
Muscovite Mica Sheets V-5 Quality	Electron Microscopy Sciences	71850-01	10 per pack
Mica Disc, 10 mm	Ted Pella, Inc	50	Mica discs are optional
Scriber Diamon Pen for Glassware	VWR International, LLC	52865-005
Scintillation Vials, Borosilicate Glass, with Screw Cap - 20 ml	VWR International, LLC	66022-060	Case of 500, with attached polypropylene cap and pulp foil liner
4 x 5 Inch Top PC-200 Hot Plate, 120 V/60 Hz	Dot Scientific, Inc.	6759-200
Straight-Sided Glass Jars, Wide Mouth	VWR International, LLC	89043-554	Case of 254, caps with pulp/vinyl liner attached
Standar-Grade Glass Beaker, 250 ml Capacity	VWR International, LLC	173506
Beakers, PTFE	VWR International, LLC	89026-022	For use with HF
Shallow form watch glass, 3"	VWR International, LLC	66112-107	Case of 12
Plastic Storage Container	VWR International, LLC	470195-354	For secondary container
General-Purpose Liquid-In-Glass Thermometers	VWR International, LLC	89095-564
High precision and ultra fine tweezers	Electron Microscopy Sciences	78310-0
Polycarbonate Faceshield	Fisher Scientific, Inc.	18-999-4542
Neoprene Apron	Fisher Scientific, Inc.	19-810-609
Calcium Gluconate, Calgonate	W.W Grainger, Inc.	13W861	Tube, 25 g
Hydrogen Peroxide 30% CR ACS 500 ml	Fisher Scientific, Inc.	H325 500	HARMFUL, TOXIC
3-Aminopropyltriethoxysilane	Gelest Inc.	SIA0610.0-25GM	Let warm to room temperature before use.
Ammonium hydroxide, 2.5 L	Fisher Scientific, Inc.	A669-212	HARMFUL, TOXIC
Hydrochloric acid	Fisher Scientific, Inc.	A144-212	HARMFUL, TOXIC
Hydrofluoric acid	Fisher Scientific, Inc.	A147-1LB	HARMFUL, TOXIC
MultiMode Nanoscope IIIa	Veeco Instruments, Inc.		Any AFM capable of tapping mode is suitable for analysis
Dunk basket	Made in lab	Made in lab	The dunk basket was made using the bottom of a PTFE bottle with holes drilled in, PTFE handle, and all PTFE screws.