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. 2025 Aug 25;5(9):4211–4223. doi: 10.1021/jacsau.5c00475

Designing Nanoparticle Surfaces with DNA Barcodes for Accurate In Vivo Quantification

Ayokunle A Lekuti †,‡,§, Vanessa Y C Li †,‡, Ayden Malekjahani †,‡, Sara Ahmed †,‡,§, Stefan M Mladjenovic †,‡,§, Marshall G G Macduff †,‡,§, Warren C W Chan †,‡,∥,⊥,*
PMCID: PMC12458034  PMID: 41001662

Abstract

DNA barcoding is a common method for identifying the biodistribution of nanoparticles. DNA barcodes are typically encapsulated within nanoparticles to ensure accurate measurements by next-generation sequencing. This method limits the types of nanoparticles that can be screened. DNA can also be coated on nanoparticle surfaces. However, it is unclear whether surface-coated DNA can be used as barcodes because they can degrade, making the identification and quantification of nanoparticle designs challenging. Here, we developed strategies to reduce DNA degradation on nanoparticle surfaces, allowing surface-based DNA barcodes for biodistribution applications. We demonstrate that nanoparticle size, DNA density, and polymer length and density are essential design parameters for accurately identifying and quantifying nanoparticles in vivo. We found that chemical modification of DNA and shielding using neutral polymers reduce DNA degradation. We validated that surface barcoding can determine the in vivo distribution of nanoparticles. Our findings pave the way for the use of surface-based DNA barcodes for in vivo screening of nanoparticle formulations for targeted applications.

Keywords: nanoparticle barcoding, DNA barcoding, surface barcoding, gold nanoparticles, DNA degradation, next generation sequencing

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Introduction

Selecting nanoparticle designs with the highest delivery efficiency to the target tissue is difficult. Mechanistic studies on the nanoparticle journey to the target tissue can lead to rational engineering for high delivery efficiency. We are still in the early stages of developing a comprehensive mechanistic framework. − A proposed alternative approach involves selecting optimal targeting formulations via nanoparticle screening. Barcodes enable the screening of thousands of nanoparticles with different permutations and chemistries. DNA sequences are the most common strategy for screening nanoparticles. − Dahlman et al. demonstrated a DNA barcoding strategy for lipid nanoparticles. The approach involves encapsulating unique oligonucleotide sequences within the nanoparticles, creating a library that can be pooled and administered to mice. After allowing the nanoparticles to circulate, the tissues are resected from the mice, and the DNA barcodes are isolated. The identities of the nanoparticle formulations within each tissue are then determined by analyzing and quantifying the DNA sequence using next-generation sequencing. Anderson et al. used this screening strategy to optimize lipid nanoparticle formulations for mRNA delivery to different tissues. Researchers have adapted this barcoding approach to identify lipid nanoparticle formulations targeting tissues in mouse models such as the liver, spleen, lungs, and tumors. −

The DNA encapsulation barcoding strategy is limited to lipid nanoparticles. This strategy involves placing the DNA barcodes in an empty nanoparticle core. Encapsulation prevents DNA degradation. This approach cannot be extended to nanoparticle designs such as gold, silica, and iron oxide nanoparticles. DNA cannot be encapsulated inside these nanoparticles. Tay et al. proposed barcoding by placing the nucleic acid sequences on the gold nanoparticle surface to investigate the delivery of differently shaped nanoparticles. Here, we examined the accuracy of quantifying DNA barcodes placed on a nanoparticle surface. We found that DNA placed on a gold nanoparticle surface was not accurately quantified. We redesigned the nanoparticle surface chemistry with various DNA densities and polymer lengths. We found that combining the chemistries minimizes degradation and provides greater accuracy in quantifying DNA barcodes at the target tissue. We demonstrated the use of surface barcoding for quantifying nanoparticle accumulation in vivo. Our strategy enables the accurate use of surface-bound DNA barcodes as identification tags to screen multiple designs in vivo. Our optimizations pave the way for engineering nanoparticle barcodes by placing DNA on their surface.

Results and Discussion

Quantifying Degradation of DNA on Nanoparticle Surfaces Ex Vivo

We began by evaluating the extent of DNA degradation on a nanoparticle surface ex vivo. DNA barcoding techniques use DNA as a proxy for quantifying nanoparticle designs. If the DNA barcodes significantly degrade, it becomes difficult to quantify and identify the nanoparticle designs accurately. The mechanism of DNA degradation is dependent on the exposure of the 5′ or 3′ termini of DNA sequences. DNA barcodes conjugated onto nanoparticles are typically conjugated with the 3′ termini exposed. DNA sequences with their 3′ termini exposed mainly breakdown via exonuclease degradation. This led us to evaluate the extent of exonuclease degradation on DNA in mouse serum and whole blood. We conjugated 79 base pair (bp) DNA sequences onto 30 nm gold nanoparticles and evaluated degradation ex vivo. We selected 79 bp DNA sequences based on the design requirements needed for probe-based quantitative PCR (qPCR) (Discussion S1). We selected probe-based qPCR to quantify the DNA sequences. We incubated the DNA-barcoded nanoparticles overnight with whole blood extracted from a mouse and varying concentrations of mouse serum. The nanoparticles were also incubated in 1× phosphate buffered saline (PBS) as a control. We quantified the DNA in the treated sample and control using qPCR. We observed that whole mouse blood and 50% mouse serum degraded DNA similarly, with only 1.5% DNA remaining (Figure S1). Over 98.5% had degraded overnight and cannot be amplified. The low DNA concentration leads to inaccurate quantification due to stochastic errors in the PCR reaction. Variable DNA length and sequence may affect the degradation rate, leading to variable quantification. − These results suggest the need to determine the design parameters for reducing DNA degradation on nanoparticle surfaces, thereby enabling surface-based strategies for accurate barcode quantification.

Polymer Shielding Reduces DNA Degradation on Gold Nanoparticle Surfaces

We first evaluated the effect of polyethylene glycol (PEG) on minimizing DNA degradation. PEG prevents serum-induced nanoparticle aggregation and opsonization. ,, We hypothesized that using PEG strands longer than the DNA length could shield the DNA from degradation. We prepared nanoparticles with oligonucleotide sequences and backfilled the remainder of the nanoparticle surface with PEG of varying molecular weights, ranging from 1 to 40 kDa, separately. Backfilling the particles with PEG had a minimal effect on the DNA density (Figure S2). We assumed that different PEG molecular weights would result in different PEG lengths on the nanoparticle surface. We used dynamic light scattering to characterize the nanoparticle hydrodynamic size (Figure S3). The hydrodynamic size increased with increasing PEG molecular weight. We incubated the DNA-conjugated nanoparticles in 50% mouse serum and 1× PBS (control) (Figure a). We chose 50% mouse serum because it degrades similarly to whole mouse blood (Figure S1). We calculated the percentage of DNA remaining on the nanoparticle surface using probe-based qPCR. Figure b and c show the percentage of DNA remaining on the nanoparticle surface coated with different PEG lengths following 4- and 24-h serum incubation. Increasing PEG length significantly reduced DNA degradation, thereby increasing the percentage of DNA remaining on the nanoparticle surface. This effect plateaued after coating the nanoparticle surface with 10 kDa PEG. We suspected that nucleases cannot fully access the DNA to cleave the sequence. To explain this effect, we measured the length of DNA and PEG. We found that the degradation plateau occurs when the PEG length exceeds the length of the DNA sequences (Discussion S2, Figure S3). These results indicated that using a PEG ligand longer than the DNA can reduce DNA degradation on a nanoparticle surface. The PEG ligand shielded the sequences from degradation enzymes.

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PEG length and density on nanoparticle surfaces affect DNA degradation. (a) Schematic workflow for evaluating DNA degradation in mouse serum and 1× PBS as a control. DNA barcoded nanoparticles with PEG from 2 kDa to 40 kDa incubated in mouse serum for (b) 4 and (c) 24 h. Statistical significance was evaluated using two-way ANOVA and corrected for multiple comparisons with the Tukey method. DNA barcoded nanoparticles with 10 kDa PEG from 0.25 to 2 PEG/nm2 incubated in mouse serum for (d) 4 and (e) 24 h. PEG is polyethylene glycol. Statistical significance was evaluated using two-way ANOVA and corrected for multiple comparisons with the Dunnett method. n = 3 replicates for each condition. All data points and error bars represent the mean ± SEM.

We investigated the effect of PEG density on reducing DNA degradation. We conjugated 30 nm gold nanoparticles with the DNA sequences. DNA-labeled nanoparticles were conjugated with 10 kDa PEG from 0.1 to 2.0 PEG/nm2. We characterized the nanoparticle sizes using dynamic light scattering (Figure S4). We incubated the nanoparticles in 50% mouse serum for 4 h. The nanoparticles were incubated in 1× PBS as a control. After the time points, the DNA sequences were quantified using qPCR. We calculated the percentage of DNA remaining for each condition. Increasing the PEG density reduced DNA degradation, thereby increasing the percentage of DNA remaining. The PEG shielding effect plateaued at a density of 0.5 PEG/nm2 and above (Figure d). This plateau effect is due to reduced DNA exposure, which occurs when the PEG molecules form brush-like structures at high densities. At low PEG densities, the PEG molecules form mushroom-like structures, thereby increasing the exposure of DNA. , PEG density controls the conformation, affecting its ability to shield the DNA sequence from nucleases. These results suggest that minimizing DNA degradation requires a PEG density greater than 0.25 PEG/nm2. We further evaluated DNA degradation after a 24-h incubation in mouse serum. We observed that the PEG density did not reduce DNA degradation. Most nanoparticle designs had the same percentage of DNA remaining (Figure e). Higher PEG densities do not reduce DNA degradation at longer time points, making DNA barcodes on nanoparticle surfaces unsuitable for long-circulating nanoparticles.

Chemical Modifications Reduce DNA Degradation on Gold Nanoparticle Surfaces

We evaluated whether chemical modifications on our DNA sequences could minimize degradation. Chemical modifications are a standard method of reducing nucleic acid degradation. ,, We examined the degradation of various modified DNA sequences (Table ). We conjugated the modified sequences onto gold nanoparticles and evaluated DNA degradation in 50% v/v mouse serum. The same nanoparticle designs were incubated in 1× PBS as a control. We calculated the percentage of DNA remaining using qPCR. Figure a shows the percentage of DNA remaining on the nanoparticle surface for each chemical modification after 4 h in mouse serum. The triple-modified sequence degraded the least and had the highest percentage of DNA remaining on the nanoparticle surface. We observed that the triple-modified sequence had 50% DNA remaining, while the unmodified sequence had only 20% remaining after a 4-h incubation. The single-modified and the 2’-O-methylation + Phosphorothioate (OME + PS) modified sequences also minimized DNA degradation, with an average of 35% remaining. The polyethylene glycol + 2’-O-methylation (PEG + OME) and PEG + PS modified sequences did not significantly reduce DNA degradation compared to the unmodified sequence. We evaluated DNA degradation after a 24-h incubation in mouse serum. We observed the chemical modifications did not reduce DNA degradation. All sequences had the same percentage of DNA remaining (Figure b). These results indicated that chemical modifications minimized DNA degradation, but only for a short time. We further investigated other ways to reduce DNA degradation.

1. List of Chemical Modifications and Their Modification Position on DNA Sequences .

Chemical modification Sequence (5′ → 3′)
Unmodified CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGGGCGCG
3′ PEG spacer CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGGGCGCG/3Sp18/
Phosphorothioate substitution CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGG*G*C*G*C*G
2’-O-methylation CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGmGmGmCmGmCG
Phosphorothioate substitution + 2’-O-methylation CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGmG*mG*mC*mG*mC*G
Phosphorothioate substitution + 3′ PEG spacer CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGG*G*C*G*C*G/3Sp18/
2’-O-methylation + 3′ PEG spacer CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGmGmGmCmGmCG/3Sp18/
Phosphorothioate substitution + 2’-O-methylation + 3′ PEG spacer CTAATACCGCATACGTCCTGAGGGAGAAAGTGGGGGATCTTCGGACCTCACGCTATCAGATGAGCCTAGGTCGmG*mG*mC*mG*mC*G/3Sp18/
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All sequences were modified with a 5′ thiol modification (/5ThioMC6-D/).

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Chemical modification and PEG lengths can reduce DNA degradation. The percentage of DNA remaining for chemically modified sequences incubated in mouse serum for (a) 4 and (b) 24 h. Chemical modifications on DNA include PEG, 2’-O-methylation, and phosphorothioate in single, double or triple combinations. Statistical significance was evaluated using one-way ANOVA and corrected for multiple comparisons with the Dunnett method. ns = not significant, **p<0.01, ***p<0.001, ****p<0.0001. Comparing the degradation of unmodified DNA (gray) to triple chemically modified DNA (pink) with PEG lengths from 2 kDa to 40 kDa on nanoparticles for (c) 4 h and (d) 24 h of incubation in mouse serum. Statistical significance was evaluated using two-way ANOVA and corrected for multiple comparisons with the Šídák method. ns = not significant, *p<0.05, **p<0.01, ****p<0.0001. n = 3 replicates for each condition. All data points and error bars represent the mean ± SEM.

We evaluated the combined effects of chemically modified sequences and PEG shielding on DNA degradation. We prepared nanoparticles with the triple-modified sequence and backfilled the nanoparticle surface with PEG of varying molecular weights, ranging from 1 to 40 kDa, separately. PEG molecules were backfilled at a density of 1 PEG/nm2. We incubated the nanoparticles in mouse serum for 4 and 24 h. The PEG molecules had a similar effect on the triple-modified sequences. The PEG length significantly increased the percentage of DNA remaining and plateaued after a 10 kDa PEG length. We compared the triple-modified sequences against the unmodified sequences. For the 4-h incubation, there was no significant difference between the unmodified and triple-modified sequences when the PEG length was greater than 10 kDa (Figure c). For the 24-h incubation, the triple-modified sequences significantly increased the percentage of DNA remaining on the surface more than the unmodified sequences, regardless of the PEG length (Figure d). These results suggest combining chemical modification of DNA and PEG length reduces DNA degradation on nanoparticle surfaces over 24 h. The chemical modifications delay the reaction of the degradation enzymes while the PEG shields the sequences from the nucleases. These alterations in the physical-chemical surface properties help reduce DNA degradation, enabling the accurate quantification of nanoparticles.

Determining the Effect of Nanoparticle Size on DNA Degradation

We investigated whether the PEG shielding strategies reduced DNA degradation for different sized nanoparticles. Nanoparticle size is an important factor in designing drug-delivering carriers. ,, Different sized nanoparticles have different surface curvatures, which we suspect will affect DNA exposure (Figure a and Discussion S3). We hypothesized that changes in nanoparticle surface curvature would decrease the PEG shielding, exposing the DNA barcodes to degradation. We synthesized nanoparticles with diameters of 18, 30, 50, and 100 nm and conjugated them with 100 DNA sequences per nanoparticle. We backfilled the DNA-labeled nanoparticles with 10 kDa PEG at 1 PEG/nm2. We incubated the conjugated nanoparticles with mouse serum for 4 and 24 h. Across nanoparticle sizes, the DNA degradation was different for the 4 and 24-h time points (Figure b–c). Differences in DNA degradation will affect the accuracy of quantifying nanoparticle designs.

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Normalizing DNA density on different nanoparticle sizes reduces DNA degradation. (a) Schematic showing the effect of nanoparticle surface curvature on DNA exposure. (b–c) DNA remaining for different nanoparticle sizes incubated in mouse serum for 4 and 24 h. Each nanoparticle had 100 DNA/nanoparticle, resulting in different DNA densities. (d) DNA remaining for 18 and 30 nm nanoparticles with varying numbers of DNA/particles after a 24-h incubation in mouse serum. Statistical significance was evaluated using one-way ANOVA and corrected for multiple comparisons with the Dunnett method. ns = not significant, **p<0.01. (e) Quantification of DNA remaining for 18 and 30 nm nanoparticles with the same DNA densities and at different concentrations incubated in mouse serum for 24 h. Statistical significance was evaluated using two-way ANOVA and corrected for multiple comparisons with the Šídák method. ns = not significant, ***p=0.0003. Quantifying the effect of DNA density on DNA remaining across all nanoparticle sizes after (f) 4- and (g) 24-h incubations in mouse serum. DNA on nanoparticle surfaces degrades similarly when DNA density is the same across all sizes. n = 3 for each condition. All data points and error bars represent the mean ± Std Dev.

Next, we evaluated if DNA density caused the difference in DNA degradation on various nanoparticle sizes. We conjugated 18 and 30 nm particles with 57 DNA and 100 sequences per nanoparticle for each size, resulting in different DNA densities (Table S1). We incubated the nanoparticles with mouse serum at the same DNA concentration for 4 and 24 h. Figure d shows the amount of DNA remaining on the surface for each nanoparticle size. The amount of DNA remaining was significantly different for different nanoparticle sizes. Whereas the amount of DNA remaining on the surface was the same within nanoparticle sizes. This result indicated that degradation was not DNA density-dependent for nanoparticles of similar size. The nanoparticle size affected the rate of DNA degradation. This result suggests quantifying different sized nanoparticles in vivo would be difficult.

We wondered if we could eliminate this nanoparticle size effect by normalizing the DNA density across nanoparticle sizes. We conjugated 18 and 30 nm with the same DNA density and incubated at approximately 700 and 1400 pM DNA in mouse serum. Figure e shows the amount of DNA remaining for each nanoparticle size. The amount of DNA remaining is the same across all conditions. Normalizing across DNA density eliminates the variable degradation. This result will allow us to accurately compare DNA barcodes for nanoparticle quantification. This result suggests that differences in DNA degradation on nanoparticle surfaces are due to differences in DNA density. To use DNA barcodes on the surfaces of different nanoparticle sizes, we need to normalize DNA density on surfaces across all sizes.

Determining the Effect of DNA Density on Degradation

We investigated whether a normalized DNA density results in consistent DNA degradation across nanoparticle sizes. We conjugated the varying nanoparticle sizes with DNA densities ranging from 0.010 to 0.20 DNA/nm2. We incubated the nanoparticles with mouse serum for 4 and 24 h. The nanoparticles were incubated in 1× PBS as a control. The percentage of DNA remaining on the nanoparticle surface was quantified using qPCR. We analyzed the DNA degradation of different nanoparticle sizes at the same DNA density. At 0.02 DNA/nm2, the amount of DNA remaining was constant across nanoparticle sizes (Figure f and g). As the DNA density increased, the percentage of DNA remaining decreased across nanoparticle sizes. These results indicated that a normalized DNA density across different nanoparticle sizes will help improve the accuracy of quantifying DNA barcodes for nanoparticle designs.

Evaluating the Accuracy and Sensitivity of Next-Generation Sequencing

We assessed the sensitivity and accuracy of next-generation sequencing (NGS) for quantifying nanoparticle concentration. We determined if DNA degradation affected nanoparticle quantification. We prepared 30 nm gold nanoparticles, separated them into four batches, and conjugated each with a unique 64 bp barcode sequence. We chose 64 bp sequences based on the design requirements for amplicon NGS (Discussion S1). We backfilled the barcoded gold nanoparticles with 10 kDa PEG at 1 PEG/nm2. PEGylation offered protection from degradation (Figure S5). We prepared a nanoparticle standard by combining the four barcoded nanoparticles in a 1000:100:10:1 concentration ratio (Figure a). The concentration of gold for each barcoded nanoparticle was quantified using inductively coupled plasma-mass spectrometry (ICP-MS) (Figure S6). We incubated the nanoparticles with 50% mouse serum for 4 and 24 h. Despite the DNA degradation, we can quantify the nanoparticles concentrations using NGS (Figure b). These results indicated that DNA degradation could be mitigated and would not significantly impact our in vivo experiments.

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Next generation sequencing (NGS) quantification of barcoded nanoparticles. (a) Schematic of workflow. Four barcoded nanoparticles were combined to create a standard curve concentration of 0.1 to 100 pM. The nanoparticles were incubated in serum for 4 and 24 h. DNA amounts were quantified with NGS, and gold nanoparticle input was quantified using ICP-MS. (b) Despite DNA degradation, NGS could quantify the relative amount of nanoparticles. Nanoparticle concentrations were relatively quantified within the sample. The NGS signal suggests that absolute nanoparticle concentration is not possible with NGS. n = 3 for each condition. All data points and error bars represent the mean ± Std Dev.

Assessing the In Vivo Effect of Surface Barcoding Nanoparticles

We investigated whether the DNA barcodes impacted nanoparticle biodistribution. If the DNA barcodes affect nanoparticle biodistribution, it would impact the accuracy and reliability of the barcoding strategy. We compared the biodistribution of DNA-labeled PEGylated gold nanoparticles to PEG-only gold nanoparticles. We conjugated 30 nm gold nanoparticles with either 64 bp DNA and backfilled their surface with PEG molecules with lengths between 1 to 40 kDa at 1 PEG/nm2. As a control, we conjugated gold nanoparticles only with PEG molecules. The diameter and charge of the particles were characterized using dynamic and electrophoretic light scattering (Figures S3 and S7). We injected BALB/c mice with each nanoparticle design via tail vein (Figure S8a). The particles circulated for 4 h, a time point at which a significant amount of DNA remains. We validated the DNA sequences are retained on the nanoparticle surface at 4 h (Figure S9). After 4 h, we euthanized the mice and resected the organs. We acid-digested the organs and quantified the gold nanoparticles using ICP-MS. The 64 bp DNA molecules significantly influenced the nanoparticle accumulation when the PEG length was shorter than 5 kDa (Figure S8b). The DNA sequence did not affect nanoparticle accumulation when the PEG length was longer than 5 kDa. The shorter PEG led to increased DNA exposure for the 64 bp sequences. We evaluated the nanoparticle surface charges and observed that the DNA-labeled nanoparticles with PEG shorter than 5 kDa had a more negative charge than the PEG-only nanoparticles (Figure S7). Sequences longer than 64 bp required significantly longer PEG (Figures S10–S11). These results indicated that the PEG length should exceed the DNA sequence to prevent DNA from influencing nanoparticle accumulation.

Validating Accurate Quantification of Barcoded Nanoparticles In Vivo

We evaluated the biodistribution of the different-sized DNA-labeled nanoparticles at the same DNA density, and PEG length and density. We synthesized nanoparticles with diameters of 18, 30, 50, and 100 nm and conjugated them with DNA sequences at 0.02 DNA/nm2 and 10 kDa PEG at 1 PEG/nm2. Nanoparticle characterizations can be found in the supplementary data (Figure S12). We administered each nanoparticle design into BALB/c mice via tail vein injections. As a control, we injected BALB/c mice with nanoparticle designs conjugated with only PEG. The particles circulated for 4 h. We euthanized the mice and resected the organs. The organs were acid-digested, and the gold nanoparticles were quantified using ICP-MS. In the blood and liver, the accumulation of DNA-labeled nanoparticles differed significantly from that of the PEG nanoparticles for both 18 and 100 nm particles (Figure a). These results were expected because the surface curvatures of the 18 and 100 nm particles differ significantly from those of the 30 and 50 nm particles. This result indicated that nanoparticles of different sizes would require surface ligands of varying lengths and densities to ensure accurate nanoparticle quantification.

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Comparing the biodistribution of DNA-nanoparticles to PEG-only nanoparticles. (a) Accumulation of 10 kDa PEG at 1 PEG/nm2 on 18, 30, 55, and 100 nm sized gold nanoparticles without DNA (gray) and with DNA (pink) in the blood, spleen and liver. (b) Accumulation of 18 nm gold nanoparticles with varying 10 kDa PEG densities without DNA (gray) and with DNA (pink) in the blood, spleen and liver. (c) Accumulation of 100 nm gold nanoparticles with varying PEG lengths and densities without DNA (gray) and with DNA (pink) in the blood, spleen and liver. Statistical significance was evaluated using two-way ANOVA and corrected for multiple comparisons with the Šídák method. ns = not significant, *p<0.05, **p<0.01, ***p<0.001, ****p<0.0001. n = 3–4 for each condition. All data points and error bars represent the mean ± Std Dev.

We focused on the 18 and 100 nm particles since they were the most inaccurate in quantification. We calculated the surface curvature of the nanoparticles (Discussion S3 and Table S1). The 18 nm particles have a higher surface curvature than the 30 nm particles. The high surface curvature of the 18 nm particles results in more space between ligands (Figure a). We hypothesized that the surface ligands are sparsely packed because of the high surface curvature of the 18 nm particles. The nanoparticles will require a higher PEG density to prevent DNA exposure. We tested our hypothesis by comparing the biodistribution of 18 nm DNA-gold nanoparticles with a PEG density of 1 or 2 PEG/nm2. We prepared the nanoparticles and injected them into BALB/c mice via the tail vein. Each nanoparticle design was administered to 4 mice. We allowed them to circulate for 4 h. We euthanized the mice, resected the organs and quantified the gold concentration accumulated in the organs. Nanoparticles with a PEG density of 2 PEG/nm2 accumulated similarly to the control, unlike the 1 PEG/nm2 coated nanoparticles (Figure b). Preventing DNA exposure for nanoparticles with high surface curvatures requires a higher density of protective ligands.

We evaluated methods to reduce DNA exposure for 100 nm nanoparticles. 100 nm particles have a lower surface curvature, resulting in surface ligands tightly packed in a brush formation (Figure a). This tightly packed formation causes increased steric hindrance. The increased steric hindrance leads to fewer ligands successfully conjugated onto the nanoparticles. Fewer surface ligands increase DNA exposure, allowing proteins to bind to the DNA barcodes. We hypothesized that 100 nm particles require longer PEG ligands at an increased density to prevent DNA exposure. We prepared 100 nm particles with DNA sequences with a 20 kDa PEG backfill to test our hypothesis. We prepared control nanoparticles with only PEG. We found the accumulation of the DNA-labeled nanoparticles was similar to that of the PEG-only nanoparticles (Figure c). Nanoparticles with low surface curvatures require a higher density and longer protective ligands to prevent DNA exposure. These results indicate that nanoparticles with low and high surface curvatures require surface ligands that are long and densely packed to avoid DNA exposure. With these designs, we minimize DNA exposure, preventing proteins from binding onto the nanoparticle and influencing its delivery.

Quantifying In Vivo Distribution of Barcoded Nanoparticles

We validated the use of surface barcoding for quantifying the in vivo distribution of nanoparticles. We prepared four different barcoded gold nanoparticles. These nanoparticles were all 30 nm in diameter, coated with 10 kDa PEG, and had a density of 1.5 PEG/nm2. Each barcoded nanoparticle was coated with a unique 64 bp sequence. These four barcoded nanoparticles were mixed at a 50:5:2.5:1 ratio (Figure S13). We injected BALB/c mice with the combined barcoded nanoparticles. The nanoparticles circulated for 4 h. After 4 h, the mice were euthanized, the organs isolated and then homogenized for NGS measurements (Figure a). The NGS analysis demonstrated that we can differentiate the ratio of the four barcodes accurately (Figure b). These measurements show the ability to quantitatively differentiate the barcodes.

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Quantifying nanoparticle accumulation using NGS. (a) Schematic of experimental workflow. Barcoded nanoparticles #1, #2, #3 and #4 were mixed in a 50:5:2.5:1 ratio. The combined barcoded nanoparticles were injected into the BALB/c mice and circulated for 4 h. Mice were euthanized, organs were resected, homogenized and sent for NGS analysis. NGS was used to quantify the DNA barcodes in the sample. (b) DNA barcodes quantified in each organ correlated with the amount of each nanoparticle injected into the mice. R 2 was calculated using a log–log line fit. n = 3–4 for each condition. All data points and error bars represent the mean ± Std Dev.

Conclusion

DNA barcoding through encapsulation is a limited technique because it cannot be used for hard or nonencapsulating nanoparticle types. Surface barcoding offers greater utility, but DNA degradation prevents the accurate quantification and identification of the nanoparticles. Optimizing the surface chemistry and design of nanoparticles can minimize degradation and enable accurate quantification. Here, we investigated chemical and physical methods to circumvent DNA degradation on surface barcodes. We reduced DNA degradation by using chemical modifications to the DNA or shielding the DNA with polymers of varying lengths and densities. We demonstrated that combining chemical modification and polymer shielding reduces the amount of DNA degraded. The DNA density should be normalized to surface area, to ensure accurate quantification of nanoparticle concentrations across different-sized nanoparticles. Our strategy enables the accurate use of surface-bound DNA barcodes as tags to screen multiple nanoparticle designs in vivo. These findings provide guidelines that pave the way for barcoding nanoparticles by placing DNA on their surface.

Materials and Methods

Custom DNA Oligonucleotides

Custom lyophilized DNA barcode sequences were purchased from Integrated DNA Technologies (Coralville, USA) with a high-performance liquid chromatography purification method. Table S2 shows details of each sequence used. The DNA barcode pellet was dissolved in Tris-ethylenediaminetetraacetic acid (EDTA, 1×) buffer. The concentration of each DNA barcode sequence was measured in a quartz cuvette using a UV–visible spectrophotometer at 260 nm absorbance. Aliquots of each sequence were prepared in 1.5 mL DNA LoBind microcentrifuge tubes (Sarstedt) and stored at −20 °C until use.

Gold Nanoparticle Synthesis

15 nm gold nanoparticles were synthesized using previously established methods. Briefly, a 250 mL Erlenmeyer flask and a 1.5″ Teflon-coated magnetic stir bar were cleaned with aqua regia (3:1, hydrochloric acid:nitric acid). The cleaned glassware was used to boil ultrapure water (100 mL). The boiling ultrapure water was rapidly stirred using the 1.5″ Teflon-coated magnetic stir bar. Chloroauric acid (1 mL of 1%w/v) was added to the boiling reaction flask. Sodium citrate (1 mL of 3% w/v, Sigma-Aldrich) was rapidly added to the boiling reaction flask. The gold chloride was reduced to ionic gold spheres in 10 min under reflux. After 10 min, the flask was transferred to an ice bath. The size distribution of the nanoparticles was measured using dynamic light scattering. Further, 30, 50, and 100 nm gold nanoparticles were synthesized overnight using the 15 nm nanoparticles as nucleation sites via the Perrault method. 15 nm gold nanoparticle solution (2.4 nM), chloroauric acid (25 mM), and sodium citrate tribasic (15 mM) were mixed in a flask. The solution was rapidly stirred with a clean 1.5″ Teflon-coated magnetic stir bar. To start the reaction, hydroquinone (25 mM) was rapidly added to the stirring solution. Table S3 contains the volumes of reagents used to synthesize 30, 50, and 100 nm gold nanoparticles. The reaction was run overnight. The next day, the reaction was quenched by adding Tween-20 (100% v/v, Sigma-Aldrich) to a final concentration of 0.05% v/v. All synthesized gold nanoparticles are concentrated by centrifugation at 3200 g (30 nm), 1200 g (50 nm), and 400 g (100 nm) in Tween-20 (0.05% v/v, Sigma-Aldrich) into 1.5 mL microcentrifuge tubes. The gold nanoparticle pellet was resuspended in sodium citrate (0.02% w/v) and Tween-20 (0.02% v/v). Nanoparticle size, polydispersity and zeta potential were characterized using Zetasizer Nano-ZS (Malvern Instruments Ltd.). Nanoparticle concentration was characterized using a UV–visible spectrophotometer (UV1601PC, Shimadzu, Kyoto, Japan). Table S4 contains the peak absorbance wavelength of 15, 30, 60, and 100 nm gold nanoparticles.

DNA and PEG Conjugation on Gold Nanoparticles

DNA barcode sequences to conjugate onto gold nanoparticle surfaces are purchased with thiol modifications on the 5′-end of the sequence,/5ThioMC6-D/. The thiol modification is reduced with tris­(2-baroxyethyl)­phosphine (TCEP) (Sigma-Aldrich) in 100× molar excess for 2 h at 37 °C on a rotator before conjugating onto the gold nanoparticle surface. Reduced thiol DNA barcode sequences are conjugated onto a gold nanoparticle surface using previously established methods. , Table S5 contains the reagent volumes for conjugating different nanoparticle sizes with DNA at 0.07 DNA/nm2. Here is a brief description of the procedure for conjugating 30 nm gold nanoparticles with DNA at 0.07 DNA/nm2. The DNA conjugation is performed in 1.5 mL DNA lobind microcentrifuge tubes. 1.95 μL of TCEP reduced DNA (10 μM) is mixed with 40 μL of sodium dodecyl sulfate (0.1% Sigma-Aldrich). The sodium dodecyl sulfate is 20% of the reaction volume. Ten μL of 18 nm gold nanoparticles (10 nM) and 105.23 μL of sodium citrate are added to the reaction mixture. The reaction mixture is vigorously mixed and incubated for 5 min at room temperature. Next, 2.83 μL of 2 kDa methyl-polyethylene glycol-thiol (5 μM, mPEG-SH, SUNBRIGHT) and 40 μL of tri-citrate-hydrochloric acid buffer (TCHB, 100 mM, pH 3.0) are quickly added to the reaction mixture. TCHB is 20% of the reaction volume. The solution is vigorously mixed and then incubated in a 60 °C water bath for 30 min. After 30 min, the DNA-nanoparticles are backfilled with PEG. The amount of PEG added varies depending on the required density. For a 1.0 PEG/nm2, 0.18 μL of 10 kDa methyl-polyethylene glycol-thiol (0.5 mM, mPEG-SH, SUNBRIGHT) is added to the reaction mixture. The reaction mixture is vigorously mixed and incubated in a 60 °C water bath for 30 min. The nanoparticles were concentrated in sodium citrate solution (0.02% w/v) by centrifugation twice to wash out the unbound DNA sequences. The pellet was resuspended in sodium citrate solution (0.02% w/v). The size distribution and concentration of the conjugated gold nanoparticles were measured using dynamic light scattering and UV–visible spectrophotometry, respectively.

Quantification of DNA Loading Density

First, the concentration of gold nanoparticles is quantified. DNA-conjugated gold nanoparticles are diluted in water to approximately 1 mL and transferred to a polystyrene cuvette for absorbance measurement on a UV–visible spectrophotometer. The peak absorbance of gold nanoparticles depends on the size of the nanoparticles. Table S4 presents the peak absorbance wavelengths of gold nanoparticles with diameters of 15, 30, 60, and 100 nm. Then, the DNA concentration is quantified. A known concentration of DNA-conjugated gold nanoparticle surface was treated with 0.1 M dithiothreitol (DTT) at 60 °C for 30 min. The thiol-DNA is displaced from the gold nanoparticle surface by DTT. The particles and displaced DNA were separated by centrifugation at 12,000g for 5 min. 0.5× SYBR gold was prepared from a concentrate in DMSO (10,000×, Sigma-Aldrich). Twenty μL of the supernatant containing displaced DNA was transferred into a chimney well black 96-well plate (Corning), and 180 μL of 0.5× SYBR gold was added to the sample. The SYBR gold intercalates into the DNA backbone. The sample’s fluorescence is measured on a plate reader (TECAN infinite 200pro) at an excitation wavelength of 496 nm and an emission wavelength of 539 nm, against a known standard curve ranging from 0 to 200 nM, which is also stained with 0.5× SYBR gold and 0.1 M DTT. The DNA concentration is calculated from the linear regression of the standard curve. DNA per gold nanoparticle is calculated using the following equation: DNA per gold nanoparticle = (Concentration of DNA)/(Concentration of nanoparticle), where the concentration of DNA is calculated from the SYBR gold assay, and the concentration of nanoparticle is the known concentration used to displace DNA for quantification.

DNA Degradation in Mouse Serum and Whole Blood

DNA-labeled gold nanoparticles were incubated in mouse serum (50% v/v with 1× PBS) (Millipore Sigma-Aldrich) or whole mouse blood at 37 °C. Mouse blood was acquired via a cardiac blood draw. Whole mouse blood was treated with 50 Units·mL–1 of heparin (BioShop) to stop blood clotting. The DNA-labeled nanoparticles were also incubated in PBS as a control. Once the incubation period ended, the degradation was quenched using a solution of sodium dodecyl sulfate (SDS), dithiothreitol (DTT) and Proteinase K. The quenching solution was added to the reaction with SDS (final concentration of 1%), DTT (final concentration of 0.1 M) and Proteinase K (final concentration of 0.5 mg/mL). The quenched solution was incubated in a 60 °C water bath for 1 h to degrade all proteins and strip the DNA sequences off the nanoparticle surface. The DNA sequences were isolated using phenol–chloroform extraction. The solution was mixed in a 1:1 volume ratio with stock phenol:chloroform:isoamyl alcohol (25:24:1, v/v, ThermoFisher Scientific). The mix was vigorously vortexed and centrifuged at 16000g for 5 min to separate the aqueous and organic phases. The aqueous layer, containing the DNA sequences, was separated from the mixture. The DNA sequences were purified from the aqueous layer using ethanol precipitation. Briefly, ammonium acetate (7.1 M), cold ethanol (100%), and stock glycogen (1 μL) were added to the aqueous layer. The ammonium acetate and ethanol were mixed in a 0.5:1 and 3:1 volume ratio with the aqueous layer. The mixed solution was incubated in dry ice (−80 °C) for 1 h. The chilled sample was centrifuged at 16,000g for 30 min to pellet the DNA. The supernatant was removed, and the sample was washed twice with cold ethanol (70%). The sample was dried using a desiccator under vacuum. The sample was resuspended using an elution buffer and quantified using qPCR.

DNA Quantification Using Probe-Based qPCR

DNA barcodes extracted and purified from tissue samples are amplified with a quantitative polymerase chain reaction. IDT DNA’s PrimeTime qPCR probe assay is used for each DNA barcode sequence amplification. Each barcode is amplified with a known concentration standard curve from 0.01 to 100 pM. The cycle threshold values for each barcode sample are calculated from concentration using the standard curve. Barcode concentrations are compared by ratios.

Elemental Analysis of Gold Nanoparticles

6 to 8-week-old BALB/c mice were ordered from Charles River Laboratory. Mice were cared for in accordance with the animal ethics protocols and oversight established by the Division of Comparative Medicine (DCM) at the University of Toronto (protocol number 20013092). A maximum of four mice were housed in cages with animal chow, automated water, housing materials, and nesting materials. To ensure mice are healthy and well-hydrated, the cages are maintained at a temperature of 21 to 24 °C and checked by DCM veterinary technician staff once or twice a week. Mice are provided with a 14/10 day and night cycle from 5 am to 7 pm and from 7 pm to 5 am, respectively. Mice were acclimated for 1 week before experimental use. Mice were injected with 100 μL of nanoparticles (150 billion particles). Mice were euthanized by isoflurane overdose, followed by cervical dislocation at 4 h postnanoparticle injection. Mouse organs were weighed and collected in 50 mL polypropylene centrifuge tubes (BioMart) for elemental analysis by ICP-MS. To collect the end point blood, the mouse’s heart was exposed, and a 25G needle connected to a 1 mL syringe was inserted into the right ventricle of the heart. The collected organs included the liver, spleen, lungs, kidneys, heart, stomach, and intestines. Nitric acid (800 μL of 16 M, Caledon 7525-1-29) and hydrochloric acid (200 μL of 12 M, Caledon 6025-1-29) were added to each organ. The samples were digested in this acid mixture for 3 h at 80 °C. Samples were diluted in deionized water (40 mL) for a final acid concentration of 2.5% v/v. For reference, a tube containing the injected dose was also digested in the acid mixture. 10 mL of all samples were filtered through a 0.22 μm PES filter (Fisher Scientific SLGP033RS) into 15 mL centrifuge tubes (Starstedt 62.554.002). A standard curve for elemental gold was prepared by diluting the elemental gold with 2% v/v nitric acid and 0.5% v/v hydrochloric acid, for concentrations from 0.0001 to 10 mg/L. All samples were quantified for gold using a NexION 350X ICP-MS (PerkinElmer) with a mass analyzer set to gold Au 197 and iridium Ir 192. A 500 μL injection loop was used, and each sample was mixed with a carrier solution (2% v/v nitric acid) and an iridium internal standard (1 μg/mL) before being injected into the analyzer. The percentage of the injected dose was calculated by the measured gold concentration of each sample divided by the measured gold concentration of the reference injected dose.

DNA Extraction from Tissue

Mice organs were collected 4 h postinjection and kept on ice. The organ tissues were chopped using a blade. Approximately 30 mg of each organ was weighed on an analytical balance (Mettler Toledo ME204TE) and used for DNA extraction. Each organ aliquot was collected in a tube (Miltenyi Biotec gentleMACS M Tube) with lysis buffer (PuroSPIN Genomic DNA Purification Plus Kit, Luna Nanotech PuroSPIN, NK261-200) and DTT (0.1 M) for homogenization (Miltenyi Biotec gentleMACS Dissociator). The organ slices were homogenized for 1 min. The tubes were centrifuged at 1,500g for 5 min to collect the homogenized organ suspension. Proteinase K (20 μL, ThermoFisher Scientific EO0491) was added to each sample and incubated in a water bath at 55 °C for 1 h. The tubes were centrifuged at 1,500g for 5 min to collect the sample. The volume was transferred into a 1.5 mL microcentrifuge tube and centrifuged at 12,000g for 3 min to pellet the denatured proteins. The supernatant (350 μL) was transferred to a new 1.5 mL microcentrifuge tube, and lysis buffer 2 (PuroSPIN Genomic DNA Purification Plus Kit, Luna Nanotech PuroSPIN, NK261-200) was added to the sample. DNA was extracted and purified using the protocol and reagents provided in the kit.

Amplicon Sequencing: Next-Generation Sequencing Prep and Analysis

DNA sequences were elongated, and partial Illumina adapters were added to the sequence. Sequence elongation was performed using two rounds of PCR amplification. All PCR reactions were performed using Q5 High-Fidelity 2× Master Mix (New England Bio). The first round of PCR consisted of 15 cycles, and the reaction used the elongation primers. The second round of PCR consisted of 25 cycles, utilizing partial Illumina adapter primers. PCR products were purified each round using the GeneJET PCR Purification Kit (Thermofisher Scientific). The PCR products were run on an agarose gel to confirm their size and purity. DNA concentration was quantified using a Qubit dsDNA Quantification Assay Kit (Invitrogen). If samples were not concentrated enough, an additional round of PCR was performed. The PCR was run for 25 cycles, using partial Illumina adapter primers. The samples are purified and quantified to ensure the quality and quantity of the amplicons. Samples were shipped to the GENEWIZ facility (Azenta Life Sciences, New Jersey, USA) for their next-generation sequencing (Amplicon EZ service) using an Illumina MiSeq platform and 250 bp paired-end reads. GENEWIZ performed the initial analysis, providing a list of all unique DNA sequences in the sample. The data was further processed to quantify DNA barcodes in the samples using a Python script. Script located at: https://github.com/AyoLekuti/Nanoparticle_surface_DNA_barcode_In_Vivo_Quantification.

Confocal Microscopy Imaging

Confocal fluorescence and reflectance microscopy was used to image nanoparticles in mice tissues. Mice were injected with 30 nm gold nanoparticles coated with 64 bp DNA and amine polyethylene glycol (aPEG). The DNA molecules were linked with Alexafluor 647 dye, and the aPEG was linked with Cyanine 3 (Cy3) dye. The gold nanoparticle core was visualized with reflectance microscopy as the gold core scatters light. The liver, spleen and kidney were harvested from the mice injected with the nanoparticles and placed into plastic cryomolds (Sakura Finetek USA, 25608-916) filled with Optimal Cutting Temperature Compound (VWR, 95057-838). The tissues were then frozen in liquid nitrogen. The frozen tissues were then cut using a Thermo Scientific HM525 NX cryostat at a thickness of 8 μm at −20 °C and adhered onto glass slides (Fisherbrand, 12-550-15). The glass slides with the tissues were then fixed in a Coplin jar for 10 min with 10% neutral-buffered formalin cat. HT501128). The slides were then washed three times for 5 min with 1× PBS in Coplin jars. The tissues on the glass slide were then traced with a hydrophobic pen. The fixed tissues were stained with Hoechst 33342 using 1 drop of NucBlue (Invitrogen, R37605) in 200 μL of 1× PBS for 30 min at room temperature. The Hoechst 333r2 stain was washed off with three washes of 1× PBS for 5 min. The tissues were then covered with 1.52 RI mounting media (Invitrogen, P36980) and coverslipped with no. 1.5 cover glasses (Fisherbrand, 12541033CA). The mounting media was cured for 24 h at room temperature while loosely covered with foil to ensure airflow. The next day, the glass slides were cleaned and imaged using a Zeiss LSM 880 inverted microscope with a 63×/1.4 NA oil-immersion objective. The brightfield images were taken with the 488 nm laser in transmitted light mode. The nuclei were imaged with a 405 nm laser. The Cy3-conjugated PEG was imaged with a 561 nm laser. The Alexafluor 647-conjugated DNA was imaged with a 633 nm laser. The nanoparticle cores were imaged with the 633 nm laser in reflectance microscopy mode with an 80% Transmittance 80% and 20% Reflectance (T80/R20) main beam splitter. The images were combined in FIJI/ImageJ.

Supplementary Material

au5c00475_si_001.pdf (1MB, pdf)

Acknowledgments

We acknowledge the Canadian Institute of Health Research (CIHR, FDN-159932; MOP-130143), Natural Sciences and Engineering Research Council of Canada (NSERC, 2015-06397), Canadian Research Chairs program (950-223824), Collaborative Health Research Program (CPG146468), and Canadian Cancer Society (705185-1) for funding support. A.A.L. and V.YC.L. would like to thank the Barbara and Frank Milligan Graduate Fellowship and Doctoral Canada Graduate Scholarship from NSERC. V.Y.C.L. would also like to thank Nanomedicine Innovation Network Graduate Award. A.M. would like to thank the Postgraduate Scholarship from NSERC. S.M.M. thanks the Ontario Graduate Scholarship, the Jennifer Dorrington Award, the NSERC Vanier Canada Graduate Scholarship and the NSERC Canada Graduate Scholarships − Master’s Scholarship. We thank Kimberly Lau and Paul Paroutis from the SickKids Imaging Facility for the use of the confocal microscope. We thank Gang Zheng, Juan Chen and James Bu at the Toronto Nanomedicine Fabrication Centre for the use of the ICP-MS instrument. We thank Atta Chang for running ICP-MS. We thank Brenda Takabe and Derek van der Kooy for the use of their cryostat. Some figures were partially created with BioRender.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/jacsau.5c00475.

  • Discussions on the DNA design, measurements of PEG and DNA lengths, and nanoparticle surface curvature calculations; supporting experimental data and details for the degradation of DNA in mouse blood and serum; DNA density of backfilled nanoparticles; size characterization of DNA- and PEG-conjugated gold nanoparticles; ICP-MS quantification of DNA-barcoded nanoparticles; charge characterization of DNA- and PEG-conjugated gold nanoparticles; influence of PEG shielding on the distribution of 149 bp DNA-conjugated gold nanoparticles; size characterization of different sized nanoparticles; ICP-MS quantification of injected nanoparticles; nanoparticle surface curvature calculations; list of DNA sequences; reagent volumes and concentrations for DNA conjugation on gold nanoparticles; and centrifugation speeds and absorbance peaks of gold nanoparticles (PDF)

#.

A.A.L. and V.Y.C.L.contributed equally to this work. A.A.L., V.Y.C.L., A.M., and W.C.W.C. conceptualized the research; A.A.L., V.Y.C.L., S.A., S.M.M., and M.G.G.M. performed the experimental research; A.A.L., V.Y.C.L., and W.C.W.C. wrote the paper. CRediT: Ayokunle A. Lekuti conceptualization, data curation, formal analysis, methodology, validation, visualization, writing - original draft, writing - review & editing; Vanessa Yen Cheng Li conceptualization, data curation, formal analysis, investigation, methodology, validation, visualization, writing - original draft; Ayden Malekjahani conceptualization, investigation, methodology; Sara Ahmed investigation, validation, visualization, writing - review & editing; Stefan Mladjenovic data curation, investigation, validation, visualization, writing - review & editing; Marshall Macduff investigation, methodology; Warren C.W. Chan conceptualization, funding acquisition, resources, supervision, writing - original draft, writing - review & editing.

The authors declare no competing financial interest.

Published as part of JACS Au special issue “DNA Nanotechnology for Optoelectronics and Biomedicine”.

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Associated Data

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Supplementary Materials

au5c00475_si_001.pdf (1MB, pdf)

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