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Selective Inactivation of Enzymes Conjugated
to Nanoparticles Using Tuned
We report a novel method for specific deactivation of conjugated enzymes using laser-
heated gold nanoparticles. Current methods involve treatment of the entire solution,
thereby inactivating all bioactive components. Our method enables inactivation of only
a single or subset of targeted enzymes. The selected enzyme is pre-conjugated to gold
nanoparticles, which are specifically heated by a laser tuned to their surface
plasmon resonance. We demonstrate inactivation of a selected enzyme, glucose oxidase,
within a mixture of biomolecules. Illumination of non-conjugated enzymes and nano-
particles demonstrated specificity. We propose a novel method to quantitatively regu-
late enzyme activity, providing a building block for cellular and cell-free biochemical
C2016 International Society for Advancement of Cytometry
enzyme inactivation; nanoparticles; laser heating
ENZYMATIC processes are widely used in a variety of disciplines such as basic
research, biotechnology, bio-engineering and diagnostics (1). Many industrial pro-
cesses also require large scale enzymatic activity. Some examples include cosmetics
(2), food production (3), textiles (4), biofuels (5), article industry (6), pharmaceuti-
cal industry (7) and more. In many processes, an enzyme needs to be active for a spe-
cific limited time period. Later on, its activity may become unnecessary or even
harmful, and thus it needs to be inactivated. A useful but still challenging task is to
inactivate an enzyme selectively without affecting other biological materials in the
medium. Current methods for rapid enzyme inactivation include dramatic pH
change or heating the entire solution above the denaturation temperature of the
enzyme (8). However, these methods lead to denaturation of all bioactive compo-
nents within the system. It is sometimes possible to inactivate a specific enzyme by
adding excess of a dominant negative enzyme or an inhibitor (8). Still, these are not
always available, and they introduce another active material into the system. Finally,
in some cases it is possible to control enzyme activity by immobilization of the
enzyme on a solid support (9,10). However, this is not applicable for applications in
which in situ localization of the enzyme is critical. We propose the ability to regulate
enzyme activity within a mixture as a fundamental building block for biochemical
reactions and therapeutics.
We have developed a new method that enables rapid inactivation of a target
protein within a solution, leaving all other molecules largely intact. Our method is
based on conjugation of the designated protein to gold nanoparticles. Conjugation
of enzymes to nanoparticles is a well-established technology that is widely used for
alterations of enzyme structure and function (9,11). Gold nanoparticles are particu-
larly suited for this purpose as they are inert and bind strongly to proteins. Gold
1Faculty of Engineering, Bar Ilan
University, Ramat-Gan 5290002, Israel
2The Bar-Ilan Institute of
Nanotechnologies & Advanced
Materials, Bar Ilan University, Ramat-
Gan 5290002, Israel
Received 21 December 2015; Revised 30
May 2016; Accepted 5 October 2016
Additional Supporting Information may be
found in the online version of this article.
*Correspondence to: Zeev Zalevsky, Fac-
ulty of Engineering, Bar Ilan University,
Ramat-Gan, 5290002, Israel. E-mail:
email@example.com and Orit Shefi, Faculty
of Engineering, Bar Ilan University,
Ramat-Gan 5290002, Israel. E-mail: orit.
A.I. and P.P. contributed equally to this
Published online 00 Month 2016 in Wiley
Online Library (wileyonlinelibrary.com)
C2016 International Society for
Advancement of Cytometry
Cytometry Part A 00A: 0000, 2016
nanoparticles are used in a large variety of biomedical process-
es such as labeling (12), drug or gene delivery (13), sensing
(14), and imaging (15,16). They are easy to produce at low
cost, chemically stable, and biocompatible (12,16–22). In case
our method will be implemented in vivo or ex vivo, gold
nanoparticles can be quantitatively detected ex vivo by atomic
absorption methods, and in vivo by CT imaging (23). Another
important characteristic of gold nanoparticles is their surface
plasmon resonance: under optical illumination, gold nanopar-
ticles efficiently create heat, which is especially significant
when the energy (wavelength) of incident photons is close to
the plasmon frequency of the nanoparticles. Importantly, the
plasmon frequency wavelength is determined by the shape
and dimensions of the nanoparticle (24,25). The heat generat-
ed by the illuminated gold nanoparticles is proportional to
the duration and energy of the illumination, and can reach
temperatures of hundreds of degrees Celsius (26). The heat is
highly localized and decreases exponentially in space, equili-
brating with surrounding temperature within a radius in the
range of a few tens of nanometers from the nanoparticle sur-
face (27–29). Thus, such heat can easily denature a conjugated
enzyme, but does not reach other proteins within the same
solution. Gold nanoparticles are commonly produced as
spheres or rods. Gold nanospheres at sizes above 2 nm diame-
ter show strong absorption at a wavelength 522 nm. Howev-
er, in gold nanorods the plasmon resonance splits into a
longitudinal mode parallel to the long axis of the rod, and a
transverse mode perpendicular to the first (24,25). As a result,
nanorods have two absorbance peaks, the primary of which
varies depending on the aspect ratio between the longitudinal
and transverse modes, and the secondary, weaker absorbance
peak always corresponds to the transverse 522 nm wavelength.
By using different shapes of gold nanoparticles conjugated to
different enzymes, selective inactivation can be achieved. The
method can be utilized for advanced biomedical applications
as well as cell-free synthetic biology applications (30,31).
Turning on (for example “hot-start” enzyme) and turning off
enzymatic activity in a quantitative manner by illumination
presents a potential method for biochemical computations
In this work we present a laser-based setup which enables
specific deactivation of a pre-selected protein, conjugated to
nanoparticles, within a mixture. We demonstrate the specific-
ity of this method to inactivate mainly the conjugated pro-
teins, by showing specific loss of activity of conjugated glucose
oxidase, in a mixture containing free horseradish peroxidase.
Gold nanospheres were purchased from Nanopartz (cat.
No. 22–30-GOAN-50 (glucose oxidase conjugated), and C11-
30-NC-50 (unconjugated). The enzymes, free and conjugated,
were from Sigma: glucose oxidase (cat. No. 49180) and horse-
radish peroxidase (cat. No. 77330). Conjugation of the gold
nanospheres to the enzyme was done by Nanopartz. Hydrogen
peroxide was from Fischer Scientific. All other reagents were
Glucose Oxidase Assay
The biochemical reaction is described in Figure 1. The
assay was performed according to the manufacturer’s instruc-
tions. 1.5 ml of 0.21 mM o-Dianisidine dihydrochloride in
50 mM KH
pH7 was mixed in a 3 ml cuvette with 300
ll of a 10% glucose solution, 4 ll of the conjugated nanopar-
ticles or free glucose oxidase (0.75U/ml), and 6 ll of a 50U/ml
peroxidase solution. Enzyme activity was measured immedi-
ately using a Nanodrop 2000c spectrophotometer at a wave-
length of 436 nm. The measurement was repeated every 5
seconds for the indicated time periods. Since we did not know
the exact number of conjugated enzyme molecules per nano-
particle, we referred to enzyme activity as the slope of the
graph of optical density vs. time.
The biochemical reaction is described in Figure 1. The
assay was performed according to the manufacturer’s instruc-
tions. 1.5 ml of a 0.21 mM o-Dianisidine dihydrochloride
solution was mixed in a 3 ml cuvette with 300 ll of double
distilled water and 4 ll of the 50 U/ml peroxidase solution.
Enzyme activity was measured immediately using a Nanodrop
2000c spectrophotometer at a wavelength of 436 nm. The
measurement was repeated every 5 seconds for the indicated
time periods. We referred to enzyme activity as the slope of
the graph of optical density vs. time.
In order to measure the activities of both glucose oxidase
and peroxidase mixed in one vial, we measured each enzyme
in half of the volume, in separate cuvettes. For glucose oxidase
activity, we added glucose, dianisidine and free peroxidase.
For peroxidase activity we added hydrogen peroxide and
An image of the laser illumination setup is shown in Fig-
ure 2. We used a continuous wave (CW) green laser 532 nm
(Photop DPGL-2100F), a 100 mm focusing lens (Newport
KPX094-C), and a folding mirror (Newport 10Z20BD.1). The
mirror diverts the beam down and thus the enzyme can be
kept in a tube, for convenience. The diffraction spot full
width half maximum (FWHM) with the focusing lens is given
where kis the laser wavelength, fis the focal length, and Dis
the beam diameter. The beam diameter was about 1 mm.
Thus, the diffraction spot FWHM was 53 lm. The average
irradiance in the focal plane depends on the diffraction spot
where Pis the optical power of the laser. In our experiments
we used the maximum output power that was 100mW, and
therefore the irradiance was 4.5 kW/cm
2Inactivation of Enzymes by Laser Illumination
The thermal measurements were conducted using a ther-
mographic camera (FLIR A325). The temperature of the solu-
tion was measured during the illumination process. Several
images from the process are presented in Figure 3A. In addi-
tion, a temperature graph with 5 sec increments is presented
in Figure 3B. The temperature data was fitted to an exponen-
tial trend-line, and found to be:
24.58C is the room temperature before we turn on the laser.
The temperature reaches a plateau at around 52.58C with a
time constant of 40 sec. A movie of the heating process is
available in the Supporting Information.
We set up a system for illumination of the enzyme-
nanoparticle solution with a 532 nm laser compatible with the
nanosphere surface plasmon resonance wavelength. The sys-
tem is described in Figure 2 and in the methods section.
To demonstrate the feasibility of the method, we focused
our efforts on commonly used enzymes, glucose oxidase and
horseradish peroxidase. Glucose oxidase catalyzes the break-
down of glucose inside cells. It has been previously shown to
remain active when bound to gold nanoparticles (34,35). Per-
oxidase functions in the roots of the horseradish plant, and
uses hydrogen peroxide to oxidize many different substrates.
Peroxidase has also been shown to remain active when bound
to gold (36). The activity assays of this enzyme combination
are well established, and can be performed in tandem as part of
a single cascaded reaction (detailed in the methods section and
Fig. 1). Thus, we could ensure that the enzymes would work
optimally in the same environment. The reaction begins with
glucose oxidase, which oxidizes glucose into gluconic acid and
hydrogen peroxide. Subsequently, peroxidase uses the hydrogen
peroxide to oxidize dianisidine, which is measured at 436 nm.
We first validated the activity assays for the enzymes using
unconjugated, free enzymes. Figure 4 demonstrates the activity
assay for free glucose oxidase, conjugated glucose oxidase, and
free peroxidase which serves as a free enzyme control in our
system. We referred to enzyme activity level as the slope of the
linear trend-line for each kinetic measurement. For Figure 4C,
the trend line slopes were 0.19 for 1 ll of enzyme, 0.34 for 2 ll,
0.46 for 3 ll, and 0.61 for 4 ll. These results indicate that the
enzyme activity assay is accurate and quantitative.
We next illuminated 4 ll conjugated glucose oxidase (dilut-
ed to 10 ll in PBS) with laser for 4 minutes, and then measured
the enzyme activity with or without illumination using the glu-
cose oxidase assay described in the methods section. Figure 5
shows that the laser illumination completely inactivated the con-
jugated glucose oxidase. This nanoparticle-based inactivation
method is similar to a common heat-inactivation. In order to
confirm that the damage to the conjugated enzyme was caused
by heat dispensed from the nanoparticles and not directly by the
laser illumination, we illuminated 4 llfreeglucoseoxidase
(diluted to 10 ll in PBS) in the absence of nanoparticles, with
no effect on enzyme activity (Fig. 5).
Figure 1. Illustration of the biochemical enzymatic assay for
activity of glucose oxidase and peroxidase. Picture was taken
from the Sigma web catalog page of the enzymatic assay of glu-
Figure 2. The system and setup. (A) An illustration of the meth-
od: the left panel illustrates the initial solution, in which gold
nanoparticles (nanospheres in this case) are conjugated to the
target enzyme (red). The solution also contains another enzyme
that is not conjugated to nanoparticles (purple). The middle panel
shows a close-up on a single nanoparticle and its surrounding.
The right panel shows that laser illumination at a wavelength
tuned to the surface plasmon resonance of the nanoparticle
(green in the case of nanospheres) heats the nanoparticle specifi-
cally, thus destroying the conjugated enzyme without damage to
the farther free enzymes. (B) An illustration of the setup. The laser
is depicted in black and the beam in green; the focusing lens is
marked by “f.” The nanoparticles-enzyme solution (red) is inside
the tube. (C) A picture of the green laser setup. [Color figure can
be viewed at wileyonlinelibrary.com]
Cytometry Part A 00A: 0000, 2016 3
nation of the nanoparticles is sufficiently localized to enable
specific inactivation of the conjugated enzyme, without damage
to the surrounding free proteins. We mixed 4 llfreeperoxidase
and 4 ll conjugated glucose oxidase (diluted to 10 llinPBS)
in the same tube, and illuminated for the indicated time peri-
ods (Fig. 6A). Our results demonstrate that the laser illumina-
tion completely inactivates glucose oxidase within
Figure 3. Thermographic measurements of the solution during the heating process. (A) Representative images at different time points.
(B) Temperature measurements average of three independent experiments taken at 5 sec increments (red) and a fitted exponential trend
line (blue). [Color figure can be viewed at wileyonlinelibrary.com]
4Inactivation of Enzymes by Laser Illumination
approximately two minutes. At that time point, 70% of the free
peroxidase is still active. Laser illumination does affect the free
peroxidase to some extent, most likely because of non-specific
heating by nearby nanoparticles. Thus, it is possible under these
conditions to eliminate the activity of conjugated glucose oxi-
dase and still keep the majority of free peroxidase active.
Figure 5. Laser illumination at 532 nm inactivates conjugated glucose oxidase specifically. 4 ll glucose oxidase free or conjugated to gold
nanoparticles were illuminated for 4 minutes or not, and activity was measured as previously described. Results are presented as means
and S.D. of three independent experiments. [Color figure can be viewed at wileyonlinelibrary.com]
Figure 4. Measurement of glucose oxidase activity. (A) Optical density values for wavelengths 350–670 nm, with each color representing
one time point in a kinetic measurement of product accumulation, every 5 seconds for 150 seconds. This panel demonstrates that the
peak absorbance was specific and as expected for oxidized dianizidine, at 436 nm. (B) Single optical density values at 436 nm, taken from
(A) for free glucose oxidase (red), specifically reflecting product accumulation. Optical density values for conjugated glucose oxidase
(blue) and free peroxidase (green) are also shown. (C) Increasing quantities of enzyme (1–4 ll) give proportionally increasing product
accumulation, i.e. enzyme activity. For convenience, OD units were multiplied by 1000 throughout the article. Results for panels (B) and
(C) are presented as means and S.D. of three independent experiments. [Color figure can be viewed at wileyonlinelibrary.com]
Cytometry Part A 00A: 0000, 2016 5
The overall temperature of the solution can provide valu-
able information to study the contribution of non-specific heat-
ing to enzyme inactivation within a solution (37). In our system,
we measured the overall temperature of the solution during the
laser illumination using a thermographic camera. The camera
allows measurement of the global temperature and not the local
temperature around the nanoparticles, however this measure-
ment provides useful information regarding the distribution of
heat inside the solution. The temperature measurement graph
and representative thermal images are presented in Figure 3. We
found that the temperature of the solution increases in an expo-
nential manner with a time constant of 40 sec, and reaches a pla-
teau at around 52.58C. This temperature is below the transition
temperature reported for glucose oxidase (38) (see discussion),
thus the inactivation of glucose oxidase in our system is directly
caused by the heat generated by the conjugated nanoparticles
and not because of the overall heating of the solution.
To further demonstrate specificity, we illuminated free
glucose oxidase in the presence of gold nanoparticles. The
nanoparticles were identical to those conjugated to glucose
oxidase, but they did not include the coating that enables
enzyme conjugation, thus leaving the enzyme soluble. The
results (Fig. 6B) show that after 1 minute of laser illumination
only 27% of the bound glucose oxidase remained active com-
pared to 73% of the free glucose oxidase.
We presented a method for denaturation and inactivation
of a specific pre-conjugated protein, in an environment that
contains other proteins that need to remain intact.
The method uses gold nanoparticles that dispense local-
ized heat in order to inactivate the conjugated proteins. We
used nanospheres and nanorods that respond to green or red
light, respectively. The optical absorption of gold nanorods
can be tailored according to needs and laser availability over a
wide range of wavelengths, from visible to near infrared
(39–41). This is especially significant for biological
applications, as near infrared light has a greater penetration
depth in biological tissues compared with visible light (39). As
reported by others (42), we also encountered the problem of
bulk heating of the entire solution by the nanoparticles, result-
ing in non-specific denaturation of proteins (Fig. 6). We dis-
covered that in order to achieve and increase specificity, a
critical aspect of the method is to adjust the illuminated area
to keep the nanoparticles under constant illumination. If they
do not match, the rapidly moving nanoparticles cannot reach
a high enough temperature to specifically denature the conju-
gated enzyme. Given enough time, if the illumination area
does not match, the nanoparticles heat the entire solution in a
non-specific manner, damaging both the conjugated and the
unconjugated proteins. The demand for diameter compatibili-
ty can be circumvented in some cases, if the designated pro-
tein has a much lower denaturation temperature than the
other proteins. Such is the case in Figure 6A, in which peroxi-
dase [with a complex temperature-dependent denaturation
curve that begins around 428C but is reversible at least up to
608C and likely up to 748C (43)] retains 90% activity after one
minute illumination in a wide tube, compared to glucose oxi-
dase [transition temperature 55.88C (38)] retains only 73%
activity after one minute illumination in a narrow tube.
In the inactivation process there is a trade-off between the
illumination fluence and the illumination duration. Stronger
illumination fluence will require less time for enzyme inactiva-
tion, but may cause too much bulk heating. Shorter illumina-
tion periods will reduce the bulk heating, but may not be
sufficient for enzyme inactivation. In addition, heating is an
exponential process. In order to achieve reproducible results, it
is preferable to work at the plateau part of the exponent. There-
fore, in our method we chose the illumination duration to be
much longer than the bulk heating time constant. This means
that the global temperature of the solution reaches its maxi-
mum, which was still below the glucose oxidase transition tem-
perature. Thus, the inactivation was caused by the local heating
generated by the conjugated nanoparticles.
Figure 6. Illumination specifically damages enzymes conjugated to nanoparticles. (A) Free peroxidase and conjugated glucose oxidase
were illuminated together in the same tube for the indicated time periods. Activity was measured as described in the methods section. (B)
Free glucose oxidase and free nanoparticles, or conjugated glucose oxidase were illuminated in separate tubes for 1 minute. Activity was
measured as described in the methods section. Glucose oxidase amount of the free enzyme was calibrated to give similar activity per vol-
ume as that of the conjugated enzyme. Results for both panels are presented as means and SEM of 4–5 independent experiments. [Color
figure can be viewed at wileyonlinelibrary.com]
6Inactivation of Enzymes by Laser Illumination
The choice of nanoparticle concentration is also impor-
tant, since a too high concentration will inevitably result in
bulk heating of the solution. The choice of nanoparticle size is
another important consideration to take into account. Pro-
teins adsorbed on smaller nanoparticles better retain their
structure and function, likely because the greater surface cur-
vature of smaller nanoparticles allows for a smaller area for
interaction between the protein and nanoparticle, and thus
lower effect on the secondary protein structure (44). On the
other hand, larger nanoparticles retain heat better, and there-
fore require shorter illumination durations and/or lower illu-
mination intensity. In our system, we eliminated the activity
of the conjugated glucose oxidase while still keeping almost
70% of the activity of the free peroxidase. A 70% level of activ-
ity is acceptable for most enzymatic applications. However, it
may be possible to further tweak and optimize the technique
in case higher activity is required, for example by modifying
the concentration of nanoparticles vs. free enzyme.
Figure 7. Illustration of example uses for our method in cytometric analyses. (A) Enzymes conjugated to gold nanoparticles are incubated
with cells. The cells uptake the nanoparticle-enzyme conjugates, and the enzymes operate within the cells. When the enzyme activity is no
longer needed, the cells are illuminated with laser at 532 nm and the enzyme is inactivated. (B) Enzymes conjugated to gold nanoparticles
and molecules that confer specificity (e.g. antibodies) are incubated with cells. The nanoparticles attach to the antigen on the cell surfaces,
and the enzymes operate on the cell. For example, here the enzyme detaches the cell from the plate and neighboring cells. Once the
desired cells are detached, the nanoparticle-enzyme conjugates are illuminated to inactivate the detaching enzyme, in order to prevent
cellular damage. The desired cells can then be collected. (C) Enzymes conjugated to gold nanoparticles, and free enzymes, are incubated
with cells. The enzymes operate in the cellular environment. When the activity of the conjugated enzyme is no longer needed, the cells are
illuminated with laser at 532 nm, and the conjugated enzyme is inactivated, leaving the other enzymes active. [Color figure can be viewed
Cytometry Part A 00A: 0000, 2016 7
Based on the advantages of the selective activation of
conjugated proteins to nanoparticles, there are many potential
applications that can be implemented to our method. Target-
ing nanoparticle complexes may affect selected populations of
cells in a highly controlled manner. Several such applications
are illustrated in Figure 7, including how our method can be
introduced into cytometric single cell analyses.
To summarize, we developed a new method for selective
enzyme inactivation that has potential applications in
research, medicine and industry. The ability to precisely and
quantitatively regulate enzyme activity may be a useful tool
for highly advanced biomedical and biotechnological applica-
tions in cellular and cell-free environments as well as future
biological computing. Our method provides means for
spatio-temporal control of enzymatic activity.
The authors thank Dr. Menachem Motiei for advice on
nanoparticles, and Shahar Levy for help with the laser setup.
1. Polaina J, MacCabe AP, editors. Industrial Enzymes. Dordrecht: Springer Nether-
2. Veit T. Biocatalysis for the Production of Cosmetic Ingredients. Eng Life Sci 2004;4:
3. Reed G, editor. Enzymes in Food Processing. New York, NY: Academic Press, Elsevier
Science; 2012. p 590.
4. Aly A, Moustafa A, Hebeish A. Bio-technological treatment of cellulosic textiles.
J Clean Prod 2004;12:697–705.
5. Zhao X, Qi F, Yuan C, Du W, Liu D. Lipase-catalyzed process for biodiesel produc-
tion: Enzyme immobilization, process simulation and optimization. Renew Sustain
Energy Rev 2015;44:182–197.
6. Bajpai P. Application of enzymes in the pulp and paper industry. Biotechnol Prog
7. Rasor JP, Voss E. Enzyme-catalyzed processes in pharmaceutical industry. Appl Catal
A Gen 2001;221:145–158.
8. Sumner JB, Somers GF. Chemistry and Methods of Enzymes. Elsevier Science; 2014. 478p.
9. Ansari SA, Husain Q. Potential applications of enzymes immobilized on/in nano
materials: A review. Biotechnol Adv 2012;30:512–523.
10. DiCosimo R, McAuliffe J, Poulose AJ, Bohlmann G. Industrial use of immobilized
enzymes. Chem Soc Rev 2013;42:6437–6474.
11. Wu Z, Zhang B, Yan B. Regulation of enzyme activity through interactions with
nanoparticles. Int J Mol Sci 2009;10:4198–4209.
12. Seekell K, Price H, Marinakos S, Wax A. Optimization of immunolabeled plasmonic
nanoparticles for cell surface receptor analysis. Methods 2012;56:310–316. Available
at: http://dx.doi.org/10.1016/j.ymeth.2011.08.017. Accessed August 28, 2013.
13. Xu L, Liu Y, Chen Z, Li W, Liu Y, Wang L, Liu Y, Wu X, Ji Y, Zhao Y, et al. Surface-
engineered gold nanorods: Promising DNA vaccine adjuvant for HIV-1 treatment.
Nano Lett 2012;12:2003–2012. Available at: http://www.ncbi.nlm.nih.gov/pubmed/
22372996. Accessed August 28, 2013.
14. Truong PL, Kim BW, Sim SJ. Rational aspect ratio and suitable antibody coverage of
gold nanorod for ultra-sensitive detection of a cancer biomarker. Lab Chip 2012;12:
1102–1109. Available at: http://www.ncbi.nlm.nih.gov/pubmed/22298159. Accessed
August 14, 2013.
15. Yang S, Ye F, Xing D. Intracellular label-free gold nanorods imaging with photoa-
coustic microscopy. Opt Exp 2012;20:10370–10375. Available at: http://www.optic-
sexpress.org/abstract.cfm?URI5oe-20-9-10370. Accessed August 28, 2013.
16. Zhan Q, Qian J, Li X, He S. A study of mesoporous silica-encapsulated gold nano-
rods as enhanced light scattering probes for cancer cell imaging . Nanotechnology
17. Hu M, Chen J, Li Z-Y, Au L, Hartland GV, Li X, Marquez M, Xia Y. Gold nanostruc-
tures: Engineering their plasmonic properties for biomedical applications. Chem Soc
Rev 2006;35:1084–1094. Available at: http://pubs.rsc.org/en/content/articlehtml/
2006/cs/b517615h. Accessed August 28, 2013.
18. Bogliotti N, Oberleitner B, Di-Cicco A, Schmidt F, Florent J-C, Semetey V. Optimiz-
ing the formation of biocompatible gold nanorods for cancer research: Functionali-
zation, stabilization and purification. J Colloid Interface Sci 2011;357:75–81.
19. Hauck TS, Ghazani AA, Chan WCW. Assessing the effect of surface chemistry on
gold nanorod uptake, toxicity, and gene expression in mammalian cells. Small 2008;
4:153–159. Available at: http://www.ncbi.nlm.nih.gov/pubmed/18081130. Accessed
August 6, 2013.
20. Rejiya CS, Kumarb J, Raji V, Vibin M, Abraham A. Laser immunotherapy with gold
nanorods causes selective killing of tumour cells. Pharmacol Res 2012;65:261–269.
21. Hu X, Gao X. Multilayer coating of gold nanorods for combined stability and bio-
compatibility. Phys Chem Chem Phys 2011;13:10028–10035.
22. Shukla R, Bansal V, Chaudhary M, Basu A, Bhonde RR, Sastry M. Biocompatibility
of gold nanoparticles and their endocytotic fate inside the cellular compartment: A
microscopic overview. Langmuir 2005;21:10644–10654.
23. Shilo M, Reuveni T, Motiei M, Popovtzer R. Nanoparticles as computed tomography
contrast agents: Current status and future perspectives. Nanomedicine (Lond) 2012;
7:257–269. Available at: http://www.ncbi.nlm.nih.gov/pubmed/22339135. Accessed
November 23, 2014.
24. El-Brolossy TA, Abdallah T, Mohamed MB, Abdallah S, Easawi K, Negm S, Talaat H.
Shape and size dependence of the surface plasmon resonance of gold nanoparticles
studied by Photoacoustic technique. Eur Phys J Spec Top 2008;153:361–364.
25. Jain PK, Lee KS, El-Sayed IH, El-Sayed MA. Calculated absorption and scattering
properties of gold nanoparticles of different size, shape, and composition: Applica-
tions in biological imaging and biomedicine. J Phys Chem B 2006;110:7238–7248.
26. Qin Z, Bischof JC. Thermophysical and biological responses of gold nanoparticle
laser heating. Chem Soc Rev 2012;41:1191–1217. Available at: http://pubs.rsc.org/en/
Content/ArticleHTML/2012/CS/C1CS15184C. Accessed July 14, 2014.
27. Govorov AO, Zhang W, Skeini T, Richardson H, Lee J, Kotov N. a. Gold nanoparticle
ensembles as heaters and actuators: Melting and collective plasmon resonances.
Nanoscale Res Lett 2006;1:84–90.
28. Avetisyan YA, Yakunin AN, Tuchin VV. Novel thermal effect at nanoshell heating by
pulsed laser irradiation: Hoop-shaped hot zone formation. J Biophotonics 2012;5:
29. Yakunin AN, Avetisyan YA, Tuchin VV. Quantification of laser local hyperthermia
induced by gold plasmonic nanoparticles. J Biomed Opt 2015;20:051030.
30. Halas NJ. Plasmonics: An emerging field fostered by nano letters. Nano Lett 2010;10:
31. Moe-Behrens GH. The biological microprocessor, or how to build a computer with
biological parts. Comput Struct Biotechnol J 2013;7:e201304003.
32. Privman V, Zavalov O, Hal
a L, Moseley F, Hal
amek J, Katz E. Networked enzy-
matic logic gates with filtering: New theoretical modeling expressions and their
experimental application. J Phys Chem B 2013;117:14928–14939.
33. Goodman JW. Introduction to Fourier optics. Roberts & Company; 2005.
34. Pandey P, Singh SP, Arya SK, Gupta V, Datta M, Singh S, Malhotra BD. Application
of thiolated gold nanoparticles for the enhancement of glucose oxidase activity.
35. Li D, He Q, Cui Y, Duan L, Li J. Immobilization of glucose oxidase onto gold nano-
particles with enhanced thermostability. Biochem Biophys Res Commun 2007;355:
36. Lin J, Qu W, Zhang S. Disposable biosensor based on enzyme immobilized on Au-
chitosan-modified indium tin oxide electrode with flow injection amperometric
analysis. Anal Biochem 2007;360:288–293.
37. Tuchina ES, Petrov PO, Kozina KV, Ratto F, Centi S, Pini R, Tuchin VV. Using gold
nanorods labelled with antibodies under the photothermal action of NIR laser radia-
tion on Staphylococcus aureus. Quantum Electron 2014;44:683–688.
38. Zoldak G, Zubrik A, Musatov A, Stupak M, Sedlak E. Irreversible thermal denatur-
ation of glucose oxidase from Aspergillus niger is the transition to the denatured
state with residual structure. J Biol Chem 2004;279:47601–47609.
39. Choi WI, Sahu A, Kim YH, Tae G. Photothermal cancer therapy and imaging based
on gold nanorods. Ann Biomed Eng 2012;40:534–546. Available at: http://www.ncbi.
nlm.nih.gov/pubmed/21887589. Accessed August 23, 2013.
40. Nikoobakht B, El-Sayed MA. Preparation and growth mechanism of gold nanorods
(NRs) using seed-mediated growth method. Chem Mater 2003;15:1957–1962.
41. Harris N, Ford MJ, Mulvaney P, Cortie MB. Tunable infrared absorption by metal
nanoparticles: The case for gold rods and shells. Gold Bull 2008;41:5–14. Available
at: http://link.springer.com/10.1007/BF03215618. Accessed August 11, 2013.
42. Della VP, Ilieva M, Zhurbenko V, Mateiu R, Faralli A, Dufva M, Hansen O. Gold
nanoparticle-based sensors activated by external radio frequency fields. Small 2015;
43. Ch attopadhy ay K, Mazumdar S. Structu ral and confo rmationa l stabilit y of
horseradish peroxidase: Effect of temperature and pH. Biochemist ry 2000;39:
44. Yap FL, Zhang Y. Protein and cell micropatterning and its integration with micro/
nanoparticles assembly. Biosens Bioelectron 2007;22:775–788.
8Inactivation of Enzymes by Laser Illumination