DEGREE / MASTER chemical... · Optical sensors for oxygen detection ..... 3 4. Optical sensors for pH ..... 5 . 1 . PRACTICAL LESSON 1: Automated portable array biosensor for multisample

OPTICAL CHEMICAL (BIO)SENSORS

GUIDE NOTES FOR LABORATORY

PROTOCOLS

DEGREE / MASTER

2016-2017

Facultad de Ciencias Químicas Universidad Complutense de Madrid

Outline

1. Automated portable array biosensor for multisample microcystin analysis in freshwater samples ............................................................................................ 2

2. Synthesis of silica nanoparticles and doping with molecular luminophores ........ 2

3. Optical sensors for oxygen detection ................................................................. 3

4. Optical sensors for pH ........................................................................................ 5

1

PRACTICAL LESSON 1:

Automated portable array biosensor for multisample microcystin analysis in freshwater samples

Automated portable array biosensor for multisample microcystin analysis in freshwater samples)

AUTOMATED PORTABLE ARRAY BIOSENSOR FOR MULTISAMPLE MICROCYSTIN ANALYSIS IN

FRESHWATER SAMPLES

Objective Development of an automated array biosensor assay based on evanescent-wave excitation for the detection of microcystins (MCs) in freshwater samples.

Note: Suggested literature for discussion in seminars previous to the laboratory practice. See reference 2: S. Herranz et al. “Automated portable array biosensor for multisample microcystin analysis in freshwater samples”. Biosens. Bioelectron. 2012, 33, 50– 55.

Introduction Microcystins (MCs) are a group of dangerous cyclic heptapeptide hepatotoxins produced by several bloom-forming cyanobacteria genera, particularly Microcystis aeruginosa spp. Cyanobacteria represent a public health concern because they can grow rapidly in waters rich in organic matter, under warm climate conditions, giving rise to the proliferation of “algae water blooms”. More than 80 MCs have been identified up to date, and microcystin-leucine-arginine (MCLR), is the most common and potent cyanotoxin. The World Health Organization (WHO) has set a guideline value of 1 µg/L for total MCLR in drinking water, the major route of human exposure. MCs promote the formation of hepatic tumors upon chronic ingestion of contaminated food or drinking water. They can also affect the kidney and the gastrointestinal tract.

MCs are commonly detected and quantified by reversed-phase liquid

chromatography (LC) combined with either ultraviolet (UV) or mass spectrometry (MS) detection after a previous sample pre-treatment step. Alternatively, biochemical screening methods, such as enzyme-linked

M. aeruginosa

MCLR


immunosorbent assays, competitive enzyme or protein phosphatase inhibition assays have also been described.

In addition, immunosensors based on different transduction schemes, such as optical (Herranz et al., 2010), electrochemical (Yu et al., 2009) and nuclear magnetic resonance measurements have been also applied to MCs analysis but they usually have limited applicability for multisample analysis.

This laboratory work is based on the covalent immobilization of MCLR onto the surface of a planar waveguide. Binding of anti-MCLR monoclonal antibodies to immobilized MCLR is competitively inhibited by the free MCLR in the sample solution. The amount of antibody bound to the patterned antigens is revealed using AlexaFluor®647-labeled rabbit anti-mouse IgG and the fluorescent signal is inversely proportional to the concentration of MCLR in the samples.

Instruments and materials Leopard Array Biosensor (Hanson Technologies, USA). The instrument

is equipped with:

ￚ A 635 nm diode laser source (LAS-635-15, Lasermax). The excitation beam is focused into the edge of the microscope slide waveguide.

ￚ The fluorescence array intensity developed on the slide is filtered using a long-pass (665 nm) and a band-pass filter (700 ± 35 nm)

ￚ Image acquisition is performed using a CCD camera (Retiga 1300, Q-Imaging).

ￚ The system is fully automated and uses two six-chamber reservoir modules, for the samples and tracers, respectively. The slide is mounted vertically and pressed against a twelve-channel gasket molded in poly(dimethylsiloxane) (PDMS, NuSil Silicone Technology), forming the six-assay flow channels. Each channel is connected on one end to a peristaltic pump and on the other to a 2-way valve that switches between the sample and the tracer that flow through the channels.

ￚ The acquired image is analyzed using a proprietary software control interface that also controls the microfluidic system. The locations and intensities of the fluorescent spots allow identifying and quantifying MCLR concentration in the samples.


(a)

(b)

Figure 1. (a) Leopard Array Biosensor (Hanson Technologies, USA), a commercial version of the NRL Array Biosensor prototype. (b) Schematic of the automated array biosensor platform used for the measurements.

Reagents:

ￚ MCLR was obtained from AbKem Iberia S.L. (Vigo, Spain). ￚ Monoclonal MC10E7 mouse IgG antibodies raised against MCs (Alexis,

Läufelfingen, Switzerland). ￚ IgG fraction monoclonal mouse anti-biotin and AlexaFluor647®-

conjugated affinity purified rabbit anti-mouse IgG were purchased from Jackson ImmunoResearch (West Grove, PA, USA).

ￚ N-hydroxysuccinimide (NHS), 2-(N-Morpholino)ethanesulfonic acid hydrate (MES)

ￚ D-biotin (Sigma-Aldrich, Mo) ￚ Poly-L-lysine solution 0.1% w/v (Sigma-Aldrich, Mo) ￚ 3-(Trimethoxysilylpropyl)diethylenetriamine (APMS)

Chip

PDMS mold

Fluidicsystem


ￚ 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) (Acros Organics, Geel, Belgium).

ￚ Tween 20 (Fluka, USA) ￚ Borosilicate microscope slides used as planar waveguides were

purchased from Thomas Scientific (Swedesboro, USA). ￚ MC stock solutions were prepared in dimethylsulfoxide (DMSO) (2

µg/mL) and stored at−20 ºC. ￚ Water was purified with a Milli-Q system (Millipore, Bedford, MA). ￚ Powder milk ￚ Alconox® ￚ All other reagents were analytical reagent grade.

Buffer assay preparation

(Use Milli-Q purified water or equivalent in all recipes and protocol steps)

MES, 50 mM, pH 6.0. Ready to use

PBS 20 mM Receipt for 250 mL (Prepare in a 250 mL borosilicate bottle) 50 mL PBS 100 mM stock solution 200 mL milli Q water

PBSTM blocking buffer: phosphate buffered saline (PBS) (20 mM, pH 7.4) containing 0.5% of Tween 20 and 3% of powdered non-fat milk:

Receipt for 30 mL (Prepare in a 50 mL falcon tube) 150 µL tween 20 0.9 g powdered non-fat milk Add 29.85 mL PBS 20 mM Mix vigorously with a vortex until non-fat milk powder is completely dissolved.

PBSTM carrier buffer: phosphate buffered saline (PBS) (20 mM, pH 7.4) containing 0.05% of Tween 20 and 0.3% of powdered non-fat milk (PBSTM, carrier buffer).

Receipt for 100 mL (Prepare in a 100 mL borosilicate bottle) 50 µL tween 20 0.3 g powdered non-fat milk Add PBS 20 mM to 100 mL


Mix with a magnetic stirrer or vortex.

Preparation of coating solutions: ￚ Poly-L-Lysine 0.01 (w/v)

Receipt for 50 mL x 2 Rinse two staining jars with 50 mL of milli-Q water, each one. Add 5 mL of poly-L-lysine 0.1% (w/v) to each staining jars. Mix gently by pipette-mixing.

ￚ MCLR and biotin coating mixtures

Reagent MCLR-mixture Biotin-mixture MCLR, 1 mg/mL 150 µL - Biotin, 2 mg/mL - 15 µL

*EDC, 800 mg/mL 37.5 µL 12.5 µL *NHS, 60 mg/mL 150 µL 50 µL

MES buffer 1162.5 µL 422.5 µL *Prepare 50 µL of EDC (800 mg/mL) and 200 µL of NHS (60 mg/mL) immediately before use.

Preparation of calibrators and immunoreagent solutions: ￚ Mycrocistis LR (MCLR, working solutions (WS))

MCLR calibrator solutions are prepared from a MCLR stock solution (1 µg/mL) following the receipt depicted in the table below.

Tube Volume (µL) of PBST

Volume (µL) of 1 µg/mL MCLR

stock

Final MCLR working solutions

concentration WS1 0 100 1 µg/mL WS2 90 10 0.1 µg/mL WS3 99 1 0.01 µg/mL WS4 99.9 0.1 0.001 µg/mL WS5 100 0 0.0 µg/mL

Capture antibodies anti-biotin y anti-MCLR (5 µg/mL) mix

Ready to use

Detection antibodies AlexaFluor® 647 anti-mouse (10 µg/mL) Ready to use


Standard solutions: MCLR standard solutions are prepared from MCLR working solutions (WS1 to WS5) prepared as depicted before. The six standard solutions necessary to perform the calibration curve are prepared as indicated in table:

Tube Volume (µL) of PBST

Volume (µL) of

MCLR WS

Capture Abs mix

(µL)

Final MCLR / “Abs mix”

concentrations*

S1 400 50 of WS1 50 100 ng/mL / 0.5 µg/mL

S2 400 50 of WS2 50 10 ng/mL / 0.5 µg/mL

S3 400 50 of WS3 50 1 ng/mL / 0.5 µg/mL

S4 400 50 of WS4 50 0.1 ng/mL / 0.5 µg/mL

S5 800 100 of WS5 100 0 ng/mL / 0.5 µg/mL

*S1 to S5 solutions should incubate at least 5 min at room temperature

Protocols Amination of the planar waveguides using poly-L-lysine

The surface was cleaned with Alconox® (Sigma-Aldrich, St. Louis, MO), intensely washed with Milli-Q water. After cleaning, the slides were immersed in a solution containing poly-L-lysine 0.01% (w/v) in water and incubated for 5 min. The aminated slides were drained and dried in 60 °C during 1 h.

Immobilization of MCLR onto the planar waveguides A 15-channel PDMS patterning gasket (see figure 2) was placed over the silanized surface of the slide, and a 150 µg/mL MCLR solution (final volume 1500 µL), containing 100 mM EDC and 50 mM NHS in MES (0.05 M, pH 6.0) was injected into each measuring channel. MES buffer and a solution containing 60 µg/mL biotin (final volume 500 µL), 100 mM EDC and 50 mM NHS were also patterned as negative and positive controls, respectively. The slides were incubated overnight at 4 ◦C and rinsed with 0.5 mL of PBS (20 mM, pH 7.4). After removal of the patterning PDMS template, the slides were incubated 2 h, at room temperature, in 3% non-fat milk in PBS containing 0.5% Tween 20 (PBSTM blocking buffer), rinsed with deionized water, dried under argon and either used immediately or stored at 4 ºC.


Figure 2. Planar waveguide functionalization using twelve-channel gasket molded in poly(dimethylsiloxane) (PDMS, NuSil Silicone Technology).

Assay protocol The measuring principle of the assay was competitive inhibition between MCLR immobilized onto the chip surface (planar waveguide) and free MCLR present in the sample for a limited number of antibody (Ab) binding sites.

Figure 3. MCLR Immunoassay Protocol: (a) Mix of samples and Abs, injection and incubation; (2) Wash the excess of bioreagents; (3) Injection of labeled detection Abs (AlexaFluor647-labeled), incubation, end washes and image acquisition. To perform the assay and, as it is depicted in figure 3, it is necessary to follow the following steps, including:

1) A volume of 400 μL of MCLR standard solution (S1 to S5) was incubated over the sensor surface for 20 min.

2) Next, the channels were rinsed (2 × 0.8 mL of PBSTM, 1 mL/min)

3) A solution containing 10 μg/mL labeled Ab in PBS was incubated for 20 min to reveal surface-bound Ab’s.

Chip de Zeonor®

Molde de PDMS


4) Unbound labeled Ab was removed by rinsing with 0.8 mL of PBSTM (4×, 1 mL/min) and the slide was then imaged.

5) Intensity data were then extracted from the charge-coupled device (CCD) images and normalized. The normalized response was plotted against the MCLR concentration on a logarithmic scale

Calculations and questions 1. Write a short resume (no more than 1,000 words) related to the

laboratory work performed: Objective, Experimental section (device description, protocols overview, format assay, etc.), Results and Discussion.

2. Indicate to which reagent corresponds the question marks of the

bioarray image data.

3. Describe concisely the mathematic models typically used to fit the calibration data in immunoassays. From them, which is the most appropriate calibration curve fitting for the work performed here? Construct a calibration plot of the data shown below and indicate on the graph the LOD and EC50.

Calibrado [MCLR], µg/L

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5

4

3

2

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202

0.38

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545

2.43

751.

565

2.37

0589

1.19

3576

2.29

451

1999

.049

1303

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-3.0

135

202.

405

280.

2255

88.5

355

190.

5405

117.

9615

186.

314

115.

239

132.

2405

180.

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0.25

815

54.3

255

1117

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50.

122

60.3

5721

31.8

86-2

1.55

6544

0.38

6594

6.85

1550

1.45

0588

3.00

149

4.47

0583

2.12

9548

5.89

3557

1.37

663

7.09

54.

2624

72.1

345

1627

.175

0.01

2955

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526

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845

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7655

1694

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511

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3833

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8865

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4175

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199

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7625

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726

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7695

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575

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7515

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5135

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0.18

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0.12

4739

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1205

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7446

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4460

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0.23

6211

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4159

4326

0.23

2923

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3919

7991

0.22

8883

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2691

5031

0.30

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670.

3031

8518

0.09

1020

210.

010.

3580

8669

0.68

3469

730.

3188

7587

0.60

3288

650.

3917

5773

0.39

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560.

4151

7846

0.40

8701

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5148

1766

0.45

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260.

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5024

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0.84

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0.52

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10.

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0.84

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References

1. “Appropriate Calibration Curve Fitting in Ligand Binding Assays”, John W. A. Findlay and Robert F. Dillard. The AAPS Journal (2007) 9, E260-E267.

2. S. Herranz, M.D. Marazuela, M.C. Moreno-Bondi, Biosens. Bioelectron. 2012, 33, 50– 55.

3. ISO, 2005. ISO Standard 20179:2005. Water Quality. Determination of Microcystins. Method Using Solid Phase Extraction (SPE) and High Performance Liquid Chromatography (HPLC) with Ultraviolet (UV) Detection. International Organization

4. “Nuevas herramientas analíticas para la determinación de contaminantes en el medio ambiente y en alimentos”, PhD Thesis S. Herranz de Andrés, Madrid, 2013.

5. S. Herranz, M. Bocková, M.D. Marazuela, J. Homola, M.C. Moreno-Bondi, Anal. Bioanal. Chem. 2010, 398, 2625–2634.

6. H.W. Yu, J. Lee, S. Kim, G.H: Nguyen, I.S: Kim, Anal. Bioanal. Chem. 2009, 394, 2173–2181.


PRACTICAL LESSON 2:

Synthesis of silica nanoparticles and doping with molecular luminophores.


SYNTHESIS OF SILICA NANOPARTICLES AND DOPING WITH MOLECULAR LUMINOPHORES.

Objective The aim of this unit is to introduce to the students to the synthesis of nanomaterials –silica nanoparticles– following a straightforward method such as the Stöber synthesis, which is based on condensation of a silica precursor in a basic alcoholic medium in the presence of controlled amounts of water. Additionally, the otherwhise colourless and transparent silica nanoparticles will be doped with a molecular luminophore, a ruthenium (II) polypyridil complex (Ru(phen)3Cl2) for providing the nanospheres with luminescent properties. Importantly, the addition timing of the Ru(II) dye in the Stöber solution has an influence on the luminescent properties of the luminescent nanobeads. The last will be evaluated by fluorescence spectroscopy.

Note: Suggested literature for discussion in seminars previous to the laboratory practice. See reference 4: D. Zhang et al. “Tuning the emission properties of Ru(phen)32+ doped silica nanoparticles by changing the addition time of the dye during the Stöber process”, Langmuir 2010, 26, 6657–6662.

Introduction The use of fluorescent nanomaterials as labels, markers or probes in bioanalysis, clinical chemistry or chemosensing is widespread. The use of fluorescence spectroscopy is of great analytical interest, since, besides a great sensitivity, this technique offers multiparametric information: emission intensity, excitation and emission wavelengths, emission lifetime and anisotropy. It is additionally, a non-destructive technique that allows remote detection and real-time / real-space monitoring.

Conventionally, molecular luminophores are used as probes or markers. Alternatively, the use of fluorescent nanomaterials offer several advantages towards the use of molecular probes such as an improved photostability and sensitivity [1]. Some luminescent nanomaterials rely in the use of the so called quantum dots, or semiconductor nanocristals, which are materials with an intrinsic luminescence with narrow emission bands whose spectral position can be tuned by modulating the QD diameter [2]. A handicap of these nanomaterials however is their toxicity, since they are normally composed of toxic elements, such as Pb or Cd. Alternatively, nanoparticles can be made of transparent and


biocompatible silica or organic polymers which have non-intrinsic luminescence, but can be doped with different types of luminophores [1].

A convenient, straightforward method for the synthesis of silica nanoparticles, is the Stöber method [2]. In this method, a silica precursor (normally tetraethyl orthosilicate, TEOS) is hydrolyzed with water in an ethanolic medium in the presence of a base (ammonia). Further condensation of the silanol groups yield spherical silica particles in the 50-2000 nm range (size particle can be tuned by modifying synthesis conditions, i.e. TEOS, water or ammonia concentration). The obtained particles can be afterwards derivatized on the surface for the introduction of functional groups that allow further bioconjugation to molecules like antibodies or DNA fragments, or conjugation of molecular fluorophres [3]. Alternativelly, molecular luminophores can be physically entrapped during Stöber synthesis into the silica network (Figure 1). Physical entrapment is feasible when using positively charged dyes that interact strongly with the negatively charged silica network. The time when the luminophore is added to the silica precursor solution has an influence on fluorophore distribution into the nanoparticles. The last, can importantly affect the luminescent properties of the doped spheres, especially in the case of the luminophore employed here, dichlorotris(1,10-phenanthroline)ruthenium(II), shortly Ru(phen)3Cl2 [4,5]. The luminescence of this metal complex is sensitive to the presence of dissolved oxygen (see Practical Lesson 3 in this Guide). The effect of addition timing of the luminophore in the emission properties of the nanospheres will be evaluated here. The dye (Ru(phen)32+) is easily adsorbed into the silica network by electrostatic interaction because of its double positive charge.

Figure 1. 1) Synthesis of silica nanoparticles following the Stöber method and, 2) addition of the luminophore Ru(phen)3Cl2 dissolved in ethanol for luminescent doping. Both processes are carried out at room temperature and with a constant magnetic stirring.

Duration: 1 session (4 h). Maximum 12 students.


Instruments and materials Centrifuge for 1.5 – 2 mL Eppendorf tubes. For nanoparticles separation

and washing.

Fluorometer for steady-state fluorescence measurements. For example, a Fluoromax-4 (Horiba Scientific) spectrofluorometer. This instrument is equipped with a 150 W xenon lamp, a photomultiplier detector and monochromator both in the excitation and the emission channels. Measurements are performed in conventional 1 cm x 1 cm cuvettes with an excitation wavelength of 450 nm (a 460 or 480 nm cut-off filter can be placed in the emission channel for decreasing the interference of scattered light). Alternatively, other laboratory spectrofluorometers can be used if they have similar characeristics (e.g. the Agilent Eclipse, Perkin-Elmer LS45 or LS55 instruments or equivalent ones from Edinburgh Instruments, Jasco, Guilden Photonics, Ocean Optics, or other manufacturers).

A stirring plate.

2.0 or 1.5 mL Eppendorf tubes.

A viewing cabinet with UV lamp (365 nm / 254 nm).

1000 µL automatic pipettes.

A magnetic bar for stirring (of the same size and shape for all students).

Disposable plastic cuvettes resistant to ethanol. Ej.: BrandTech® Macro Fluorescence Cuvettes for UV (1 cm x 1 cm, V = 2.5 – 4.5 mL); or Kartell (1 cm x 1 cm, 4.5 mL, 4 clar faces, art. Nº 1961, toghether with LDPE 1cm x 1cm stoppers: http://www.kartelllabware.com/en/products/dispolab/cups-and-cuvettes/uv-range-cuvettes/)

Reagents: ￚ Tetraethylorthosilicate (TEOS, 99%). ￚ Ammonium hydroxide (aqueous solution, 28% w/w or 16.5 M). ￚ Dichlorotris(1,10-phenanthroline)ruthenium(II), CAS: 207802-45-7 (for

example, from Aldrich, Art. Nr.: 343714). A Stock solution (c = 0.2 mg/mL) of this fluorophore will be provided in absolute ethanol.

ￚ Absolute ethanol. ￚ Type 1 ultrapure water (Milli-Q). ￚ A facility for purging the solutions of the fluorescence cuvettes with an inert gas (preferable argon; nitrogen could also be used).


Protocols Synthesis of silica NPs doped with Ru(phen)3Cl2.

Silica nanoparticles containing Ru(phen)32+ 2Cl- are prepared according to the Stöber method. Typically, 4 mL of ethanol are mixed with 235 µL of water and 235 µL of 28% ammonia in a 10 mL glass vial with a stirring bar and protected from light. The final ethanol solution (2.9 M in water and 0.9 M in ammonia) is stirred at a constant speed of 150 rpm at 25 ºC and the reaction is started with the addition of 335 µL of TEOS (1.5 mmol). This is the starting time of the reaction, t0 = 0 min. After a certain time (3 min for sample NP1, 1:30 h for sample NP2 or 2:20 h, for sample NP3), 335 µL of a 0.20 mg/mL solution of Ru(phen)3Cl2 (c = 0.24 µmol/mL) in absolute ethanol is added. The resulting solution is stirred for 2:25 h (overall time since TEOS addition), and splitted into 3 Eppendorf vials (1500 µL in each Eppendorf). The nanoparticles are isolated by centrifugation (6 min, 11000 rpm). The yellow solid remaining at the bottom of each vial (see Fig. 2) is re-dispersed in 1500 µL of fresh ethanol, and the suspension is centrifuged again. The remaining yellow solids at the bottom of each 3 Eppendorf are joined in a single Eppendorf and re-dispersed and stored in 1500 µL of fresh EtOH. The emission of the corresponding suspension (sample NP1, NP2 or NP3) can be directly observed with the naked eye under UV irradiation. For this, add 500 µL of the corresponding NP1, NP2 or NP3 sample in a small glass vial and add 500 µL of absolute ethanol for each sample. Place the vials under UV light (365 nm) in a viewing cabinet and compare the emission intensity of each NPs sample.

Figure 2. The luminescent nanoparticles are separated by centrifugation and washed with fresh ethanol. The luminescence can be observed by the nacked-eye under UV-light excitation (365 nm). A transmission electron microscopy (TEM) image shows that the obtained nanoparticles have a spherical shape and a diameter of ca. 200 nm.

Spectroscopic characterization of the Ru(phen)3Cl2 luminohore and

the luminescent NPs. 1) Emission properties of the luminescent complex Ru(phen)3Cl2 in ethanol. While waiting for luminophore addition to NP2 or NP3 at time 1:30 h or 2:20 h, respectively, the molecular luminophore Ru(phen)3Cl2 will be


characterized by fluorescence spectroscopy in ethanol solution under air, and, after bubbling argon for 10 min. For this, 1 cm x 1 cm plastic cuvettes are filled out with 2.5 mL of absolute ethanol. Then 110 µL of the Ru(phen)32+ Stock solution are added (cf = 10-5 M). Emission is recorded between 520 and 850 nm by exciting at 450 nm (integration time 0.2 s, step: 1 nm). A 460 or 480 nm optical cut-off filter can be placed in the emission channel for decreasing interference from the scattered light. After this recording, the same cuvette, sealed with a rubber septum or a plastic stopper with parafilm, is purged with argon for 10 min –with the aid of a needle connected to the argon source– to eliminate dissolved oxygen. Immediately after, place the cuvette in the spectrofluorometer and record the emission (same conditions as before). Especial care must be taken in avoiding oxygen diffusion inside the cuvette after the argon purge. 2) Emission properties of fluorescent nanoparticles. 1 cm x 1 cm plastic cuvettes are filled out with 2.5 mL of absolute ethanol. Then, 100 µL of the corresponding NP suspension (NP1, NP2 or NP3) are added. The emission is recorded with the same experimental conditions as for the Ru(phen)3Cl2 dye (always shake the NPs suspension in the cuvette immediately before recording the emission, otherwise the NPs can deposit at the bottom). As in the previous part, the cuvettes with the NPs suspensions are sealed with a septum and purged with argon for 10 min. Emission is again collected immediately after argon bubbling and compared with the results obtained before the argon purge.

Questions 1. What happens to the emission intensity of the luminophore Ru(phen)32+ in

ethanol solution after bubbling argon for 10 minutes and why? 2. Compare the results obtained by fluorescence spectroscopy for the three

samples of doped nanoparticles: NP1, NP2 and NP3. Which sample has a stronger emission? Which sample is more sensitive to the presence of dissolved O2? Justify the observed results according to ref. [4].

3. An alternative to the Stöber method for obtaining monodisperse silica NPs is the microemulsion method (see ref. [6]). Describe its synthetic principle and comment the differences with respect to the Stöber synthesis.

References

1. S. Bonacchi, D. Genovese, R. Juris, M. Montalti, L. Prodi, E. Rampazzo, M. Sgarzi, N. Zaccheroni “Luminescent chemosensors based on silica nanoparticles”, Top Curr Chem 2011, 300, 93–138.


2. Original work from Söber: W. Stöber, A. Fink, E.J. Bohn, “Controlled growth of monodisperse silica spheres in the micron size range”, Colloid Interface Sci. 1968, 26, 62–69.

3. S. Won Bae, W. Tan, J.-I. Hong, “Fluorescent dye-doped silica nanoparticles: new tools for bioapplications”, Chem. Commun. 2012, 48, 2270–2282.

4. D. Zhang, Z. Wu, J. Xu, J. Liang, J. Li, W. Yang, “Tuning the emission properties of Ru(phen)32+ doped silica nanoparticles by changing the addition time of the dye during the Stöber process”, Langmuir 2010, 26, 6657–6662.

5. A.B. Descalzo, C. Somoza, M.C. Moreno-Bondi, G. Orellana, “Luminescent core−shell imprinted nanoparticles engineered for targeted Förster resonance energy transfer-based sensing” Anal. Chem. 2013, 85, 5316−5320.

6. X. Zhao, R.B. Bagwe, W. Tan, “Developmet of organic-dye-doped silica nanoparticles in a reverse microemulsion”, Adv. Mater. 2004, 16, 173−176; K.S. Finnie, J.R. Bartlett, C.J.A. Barbé, L. Kong, “Formation of Silica Nanoparticles in Microemulsions”, Langmuir 2007, 23, 3017−3024.


PRACTICAL LESSON 3:

Luminescent sensors for oxygen measurements


LUMINESCENT SENSORS FOR OXYGEN MEASUREMENTS

Objective Fabrication and testing of an optical sensing film for molecular oxygen measurements in air or in water.

Note: Suggested literature for discussion in seminars previous to the laboratory practice. 1) G. Orellana, et al. “Oxygen Sensing in Nonaqueous Media Using Porous Glass with Covalently Bound Luminescent Ru(II) Complexes”. Anal. Chem. 1998, 70, 5184–5189 and, 2) G. Orellana et al. “Oxygen-Sensitive Layers for Optical Fibre Devices”. Mikrochim. Acta 1995, 121,107–118.

Introduction Sensors, both physical or chemical ones depending on their stimuli, are

regarded currently as the “senses of electronics” because they allow reception of inputs from the environment of the latter (Figure 1). Optical sensors (sometimes termed “optodes” to differentiate them from their electrical, thermal, acoustic or mechanical counterparts) have reached a leading position in those areas where the features of light and of the light-matter interaction show their superiority: contactless and/or long distance monitoring, detection sensitivity, wavelength selectivity, absence of electrical interference or risks and lack of analyte consumption, to name a few [1]. The introduction of optical fibers and integrated optics has added value because they can confine and readily carry light to inaccessible locations, higher information density may be transported, specific indicator dyes can be used for unique (bio)chemical sensing, and optical sensors can be mass produced, while additional optosensing schemes have been established (interferometric, surface plasmon resonance, energy transfer, supramolecular recognition,...).


Luminescent chemical sensors (i.e. those based on the emission of light from the lowest-lying electronic excited state of particular molecules,1 typically upon illumination with a source of shorter wavelength than the observed luminescence [2]) occupy a prominent place among the optical devices [1]. This is due to its superb sensitivity (just a single photon sometimes suffices for quantifying emission), combined with the required selectivity that electronic photoexcitation imparts to these devices.

Figure 1. General scheme of a chemical sensor depicting its essential components. The specific analyte-sensitive indicator dye embedded into a polymer film (green area) imparts selectivity to the sensor. If an additional biological component is used to enhance the device selectivity (white recognition sites), a chemical “biosensor” is produced. Interaction of the target analyte with the indicator dye elicits or modulates the optical response, which is quantified by a photodetector. Other transduction principles are the basis of alternative chemosensors.

1 The emission of light from the lowest-lying electronic excited states of molecules is known as

“fluorescence”, if arises from a “singlet” excited state, or “phosphorescence”, if the emitting excited state is of a “triplet” type. A more detailed information can be found in [2].


Figure 2. Scheme of a luminescent chemical sensor for molecular oxygen (O2) measurements in water or in air. The analyte-sensitive indicator-doped polymer film is commonly immobilized at the distal end of an optical fiber (center). Upon excitation with blue light (nowadays a LED source) travelling through the fiber, the red luminescence from the sensitive terminal has maximum intensity in the absence of the analyte (bottom left). However, the presence of O2 in the gas-permeable sensor film quenches the dye luminescence in a proportional way to its concentration in the fluid. The variation of the luminescence intensity is quantified by a portable luminometer. The real-time data collected by the luminometer can be (remotely) downloaded to a computer for environmental and industrial monitoring of O2.

We will illustrate this lab experiment with an example of luminescent sensor that has jumped the gap that separates laboratory research from the instrumentation marketplace. The use of sensors avoids the “traditional” (slow, costly) way of analyzing water or air by manual sampling, transportation to a chemical laboratory and determination of the analyte of interest by dedicated personal. Remote sensing allows in situ, real-time, continuous field monitoring of chemical species, effectively sending the lab-to-the-sample instead of the sample-to-the-lab [3].

Luminescent optical sensors can measure either the intrinsic luminescence of the analyte (very few examples known) or design sensors based on the variation of the luminescence of an indicator dye with the determinand concentration. The probe molecule must be immobilized onto a polymer support (sometimes the waveguide itself) and placed at the distal end or in the evanescent field of an optical fiber or integrated optics sensing device (e.g. see Figure 2).

The (electronic) excited state of the indicator dye (D*) undergoes a so-called photochemical reaction with the analyte (O2). Photochemical reactions are uni- or bi-molecular chemical processes induced by light that involve an excited state of an atom or molecule generated by absorption of photons (hν). Therefore, light is not a catalyst, but it is consumed in the course of the photochemical reaction:

2

D

O22

D

O2


D + hν → D* D* (+ Q) → products Bimolecular photochemical reactions are typically of the electron transfer

(“photoredox”), proton transfer (“photoacididy/photobasicity”) or energy transfer (“photosensitization”) type. While the first two types have their equivalent in regular (ground-state) chemical processes, the last one has no ground-state analogue as it involves a transference of the energy of the excited state of the photon-absorbing molecule to the reaction partner, subject to specific rules [4]. In the case of the indicator dye (abbreviated RD3, see below) excited state “quenching” by O2 of this practical lesson, the photochemical reaction leads to the ground state of the former and an electronic excited state of O2 called “singlet” (molecular) oxygen (*O2). The latter returns to its ground state in a few microseconds so that, unlike electrochemical O2 sensors [3], there is no oxygen consumption:

RD3 + hν → *RD3 *RD3 + O2 → RD3 + *O2 *O2 → O2

The variation of the indicator dye luminescence intensity with the O2 concentration in the indicator solution (or with the O2 partial pressure, PO2 in the case of gas phase measurements) obeys a linear relationship called the “Stern-Volmer equation” [2,4]:

𝐼𝐼0𝐼𝐼 = 1 + 𝐾𝐾𝑆𝑆𝑆𝑆[O2]

where I0 and I are the luminescence intensity of the indicator dye (or the sensor film) in the absence and in the presence of a particular O2 concentration (or partial pressure).

Modern optical sensors for O2 measurements have found application in environmental monitoring (water courses and reservoirs, oceans, landfills of waste, sediments), industrial analysis (chemical processes, petrochemical, biogas/biomethane production monitoring, biofuel production, nuclear power plants refrigeration water, storage of flammable chemicals,…), aeronautics (in-flight inertization of fuel tanks), biotechnology (bioprocess and bioreactor monitoring), medicine (in vivo O2 monitoring in tissues and organs), biology (intracellular measurements of O2 levels), among other areas. due to their robustness, high sensitivity, fast response, capability of being miniaturized, competitive price, low maintenance, and absence of electrical interferences or risks.

This laboratory work aims to illustrate the principle of indicator-based luminescence optosensing of dissolved O2 by observing the effect of this gas on the luminescence of a tailored indicator dye in solution. Then, we will manufacture an indicator-doped polymer thin film identical to those used in commercial optical sensors for O2 monitoring and we will test its response to the analyte species in the gas phase.


Instruments and materials UV lamp for TLC plates or gels visualization (e.g. dual-wavelength

365/254 Vilbert Lourmat™ UV lamp model VL-6.LC, Germany; http://www.vilber.de/en/products/uv-instruments/uv-lamps/filtered-uv-lamps/). (Figure 3).

Figure 3. Dual-wavelength (365/254 nm) Vilbert Lourmat™ UV lamp model VL-6.LC. Single-

wavelength (365 nm) models of any length can also be used for this experience.

Reagents:

ￚ Tris(4,7-diphenyl-1,10-phenanthroline)ruthenium(II) dichloride, abbreviated

RD3 (Sigma-Aldrich, now Merck, ref. 76886, Germany) (Figure 4) [5]. ￚ Chloroform, stabilized with ethanol, for analysis ACS (PanReac

AppliChem, ref. 131252, Spain). ￚ Neutral-cure RTV silicone (Quilosa, N-26 translucid, Spain) (or any other

1-part translucid one of those used for bathroom or windows sealing). ￚ 76 x 26 mm microscope glass slides, cut edges (VWR International

Eurolab, ref. 631-1550, Spain) ￚ Polyester film (e.g. A4-size overhead transparent sheets for laser printer

or copier, or for manual drawing) ￚ Polyester adhesive film 70 µm (Delex, Spain) (any thickness between 70

and 120 µm will be fine) ￚ Laboratory tweezers for microscope slides (Los Productos de Aldo, ref.

PD1420, Spain) ￚ Glass vials, 10 mL, 23 mm ∅, PE lid (Los Productos de Aldo, ref.

VNC23046, Spain) ￚ Disposable non-sterile LDPE 1.5-mL graduated transfer pipettes (e.g.

VWR International Eurolab, ref. 612-3399) ￚ Argon canister (LAB-10, Contse, Spain) with pressure regulator, silicone

tubing and plastic pipette tip (for 100-mL pipettes) fitted to the distal end of the tubing. If argon is not available, pure nitrogen gas can be used instead.

http://www.vilber.de/en/products/uv-instruments/uv-lamps/filtered-uv-lamps/


Figure 4. Chemical structure of the tris(4,7-diphenyl-1,10-phenanthroline)ruthenium(II) dichloride, abbreviated RD3, red-luminescent O2 indicator dye used in this experience. For the

sake of clarity, hydrogen atoms are not shown in the 3-D drawings.

TIP: If a laboratory spectrofluorometer is available to the students (e.g. the Agilent Eclipse or Perkin-Elmer LS45 or LS55 instruments or equivalent ones from Edinburgh Instruments, Horiba, Jasco, Guilden Photonics, Ocean Optics, or other manufacturers), this experience may be significantly enriched by measuring the excitation and the luminescence spectra of the RD3 indicator dye in solution and the emission intensity of both the solution and the sensor membrane under air and under pure argon or nitrogen (while being flushed into the fluorescence cell containing either the solution or the dye-doped silicone film, placed in the sample holder of the instrument). The “kinetic” mode of the spectrofluorometer may be selected to have luminescence measurements at the emission wavelength maximum in real time, so that the O2-sensitive film response and its sensitivity (IAr/Iair) can be measured for different films prepared by the students, or at different dyeing times or film thicknesses.

Indicator-doped film preparation

The quantities specified below are enough for 15 students; the chloroform solutions should be better prepared under a laboratory hood. Reproduction of the video clips requires a live internet connection.

Indicator dye solution in chloroform.

Prepare 20 mL of a ca. 0.5 mg/mL solution of RD3 in chloroform and distribute it in 5 glass vials. IMPORTANT: This solution is not exhausted in the dyeing process; it may be used for doping more than 50 membranes/vial if properly kept protected from light and tightly closed in


the container, preferably in the fridge (the solution is non-flamable). Using a permanent marker (e.g. Staedtler Lumocolor), make a mark of the liquid level on the glass vial before storage; if solvent has evaporated, add chloroform up to the mark before further use.

Silicone film.

Cut approx. 60 x 30 mm rectangles of the overhead sheets (polyester film) (2−3 per student)

Cut approx. 5 mm-wide stripes of the polyester adhesive film and stick two of them onto the edges of each polyester rectangle. Remove the stripe exceeding the rectangle length:

Place the cast mold onto a paper sheet and deposit a small amount of the RTV silicone onto it (between the plastic stripes), leaving ca. ¼ of the cast length in one side and ca. ¾ of its length onto the other:

With the aid of a microscope slide sitting onto the plastic stripes, spread the silicone onto the polyester cast mold pressing firmly against the underlying plastic stripes. Hold firmly the horizontal cast against the lab bench. Try with a few slides until you get a uniform silicone film. Do not worry if the excess of silicone overflows the end of the mold. Allow the manufactured silicone film to cure at room temperature for 72 h under a relative humidity higher than 30%. A higher RH level and/or temperature speeds up the curing process. The film is tackle-free after just 60 min under RH > 30%; however, optimum physical properties are not reached until 72 h of curing. Therefore, the instructor should manufacture 15-20 preparations before the practical session in order to continue with the students work flow. The current student preparations may be used for the following group of students because the cured silicone films are stable for several years.

With the help of the laboratory tweezers, detach the cured films from the underlying plastic mold and leave them on a clean sheet of paper. In this


moment, they are ready for the indicator dye doping. The aspect of the uncured and cured silicones films can be visualized in Video 1 below.

Video 1. Appearance of a just-spread and a cured silicone film.

Doping the silicone films with the indicator dye.

Hold the cured silicone film with the lab tweezers and dip it into the dye solution. Keep it into the solution for 5−8 min, moving it occasionally with the help of the tweezers or shaking the container gently. Remove it from the solution and deposit it onto the paper sheet. Observe what happens while the solvent evaporates. Allow a few minutes for the solvent to completely evaporate. The process of dyeing the silicone film can be visualized in Video 2 below.

Video 2. Dyeing of a cured silicone film with the luminescent indicator.

The appearance of several indicator-doped films prepared by the students themselves can be observed in Video 3 below.

https://www.youtube.com/watch?v=Sy7_9aCuhHM

https://www.youtube.com/watch?v=NFqbTmCD21E


Video 3. Appearance of indicator-loaded silicone films for luminescence O2 sensing.

Experiments The results are more striking if carried out under dimmed lighting conditions.

Effect of dissolved O2 on the luminescence of the indicator dye

Use a hood to carry out this experiment. Place 1.5 mL of chloroform in a glass vial with a LDPE transfer pipette and add 0.1 mL of the indicator dye solution (see above) with the same pipette. Prepare two identical solutions with the same dilution, each into a different vial, and close them with their caps. Place the vials containing the solutions on top of the UV lamp and switch on the 365 nm source. Observe the luminescence coming from the solutions.

Open one of the vials (or make a small hole in its flexible plastic lid so that the plastic pipette tip can go through), and strip out the dissolved air by gentle flushing the solution with an argon stream during ca. 5 min while observing simultaneously the luminescence from the closed vial and from the one being sparged with argon (Video 4).

Video 4. Luminescence from the indicator dye solution under air and after sparging with argon, upon excitation at 365 nm.

https://www.youtube.com/watch?v=jjoyp-wMmOc

https://www.youtube.com/watch?v=mKojKLWMnIM


Effect of O2 on the luminescence of the indicator silicone film

Place the indicator-loaded silicone film onto the UV lamp and switch on the 365 nm source. Observe the luminescence coming from the sensor film. Approach the plastic tip of the tubing connected to the argon canister to the O2 sensor film and observe the effect on the luminescence coming from the region of the film hit by the argon stream. Sometimes the effect is easier to visualize if two sensor films are sandwiched and placed onto the UV lamp.

Video 5. Effect of O2 on the luminescence of a sensor film loaded with indicator dye. In this case, two sandwiched sensor films have been used for a more effective visualization.

https://www.youtube.com/watch?v=ig71DEjeIvE


Questions 1. Explain why the indicator dye luminescence (both in solution and

embedded in the polymer film) increases when O2 is removed from the solution or from the vicinity of the polymer film. Note: Do not use the Stern-Volmer equation to explain it! The equation just provides a quantitation of the oxygen effect…

2. (a) Why the dyed membrane shrinks when removed from the chloroform

solution? (b) Why its color is different on the paper sheet and when illuminated with the UV lamp? (c) Why do we illuminate it with UV light instead of blue light, which is also strongly absorbed by the (orange) indicator dye? (d) If we use a blue LED for exciting the solution/sensor luminescence, what would we need to observe the red luminescence from the indicator dye with our eyes?

3. Figure 5 depicts the dose-response curve of an actual luminescent O2

sensor to the analyte variations in the gas phase. If the sensor is introduced in water and the dissolved O2 concentration is varied by flushing the solution with O2/N2 mixtures of the same concentrations, the sensor response intensity is identical (but not its response time). Explain why the sensor response is identical in solution and in the gas phase.

Figure 5. Typical dyed silicone film used for luminescence-based O2 measurements from which sensor disks are cut [6]. The graph on the right side depicts actual oxygen measurements in the gas phase of O2/N2 mixtures at atmospheric pressure. A detailed study on the reasons for the non-linear response of the silicone-based sensor films can be found in [7].


References

1. G. Orellana, “Fluorescence-based sensors”. In: Optical Chemical Sensors, F. Baldini, A.N. Chester, J. Homola, S. Martellucci, Eds. NATO Sci. Ser. II, Vol. 224., Springer-Kluwer, Amsterdam, 2006; pp. 99-116.

2. B. Valeur and M.N. Berberan-Santos, Molecular fluorescence: Principles and Applications, 2nd ed.; Wiley-VCH, Weinheim, Germany, 2012.

3. G. Orellana, C. Cano-Raya, J. López-Gejo, A.R. Santos, “Online Monitoring Sensors”. In: Treatise on Water Science, Vol. 3, P. Wilderer (Ed.), Elsevier, Oxford, UK, 2011; pp. 221-262.

4. V. Balzani, P. Ceroni, A. Juris, Photochemistry and Photophysics: Concepts, Research, Applications, Wiley-VCH, Weinheim, Germany, 2014.

5. G. Orellana and D. García-Fresnadillo, “Environmental and Industrial Optosensing with Tailored Luminescent Ru(II) Polypyridyl Complexes”. In: Optical Sensors: Industrial, Environmental and Diagnostic Applications, R. Narayanaswamy and O.S. Wolfbeis (Eds.), Springer, Berlin-Heidelberg, Germany, 2004; Ch. 13, pp. 309-357.

6. G. Orellana, J. López-Gejo, B. Pedras, Silicone films for fiber - optic chemical sensing. In: Concise encyclopedia of high performance silicones, Wiley, Hoboken, New Jersey, 2014, pp. 339-353.

7. J. López-Gejo, D. Haigh, G. Orellana, “Relationship between the microscopic and macroscopic world in optical oxygen sensing: A luminescence lifetime microscopy study”, Langmuir 2010, 26, 2144-2150.


PRACTICAL LESSON 4:

Determination of pH in aqueous samples by fluorescence using a phase-sensitive sensor


DETERMINATION OF pH IN AQUEOUS SAMPLES BY FLUORESCENCE USING A PHASE-SENSITIVE SENSOR

Objective Development of an automated optical sensor based on phase shift measurements for the determination of pH in river waters.

Note: Suggested literature for discussion in seminars previous to the laboratory practice. See reference 3: W. J. Bowyer et al. “Determining Proton Diffusion in Polymer Films by Lifetimes of Luminescent Complexes Measured in the Frequency Domain”. Anal. Chem. 2009, 81, 378–384; H.M.R. Goncalves et al. “Fiber optic lifetime pH sensing based on ruthenium(II) complexes with dicarboxybipyridine”. Anal. Chim. Acta, 2008, 626, 62–70.

Introduction The pH is a chemical parameter whose –sometimes quick– strict and continuous control is essential in many chemical processes in the food, pharmaceutical as well as in the medical fields. In most of these areas, the measurement of the pH is performed using a conventional glass electrode in which the pH reading is produced as a result of an exchange of hydrogen ions between the sample and the hydrated gel layer which coats the inner and outer surfaces of the glass. Despite the glass electrode provides fast, reliable and accurate measurements, it is not always feasible to use this method for determining the pH. In recent decades, fluorescence studies relying in the measurement of the emission lifetime (τ) of a luminophore have proliferated, because they provide more information than methods relying in steady-state fluorescence measurements. There are two techniques for determining the emission lifetime of a luminescent indicator: i) time-resolved detection and ii) phase sensitive detection. i) In time-resolved measurements the luminophore is excited with a pulse of light, then the measurement is the extinction of the luminescence of the compound function of time. The emission lifetime is thus calculated from the kinetics of deactivation from the excited state of the indicator, which corresponds to the inverse of the rate constant with which the unimolecular kinetic process of the photoexcited luminophore deactivation takes place. I(t)= Io exp(−t /τ )


ii) The second method used to determine emission lifetimes is based in phase-sensitive detection. In contrast to time-resolved measurements, the sample is excited with sinusoidally modulated continuous light, so that the emission of the probe is modulated to produce the same excitation frequency, but out of phase with it due to the time that the molecule remains in the excited state.

Figure 1. Schematic representation of the phase (φ) and demodulation (m) of the emission of a compound with respect to the excitation light.

Figure 1 shows that not only does a gap (φ) occur in the light emission from the luminophore, but also the modulation factor is reduced (m, amplitude ratio) with respect to the excitation. From these two parameters, when the deactivation kinetics is strictly mono-exponential, it is possible to determine the τ luminophore according to equations 1 and 2. 𝜏𝜏∅ = tan∅

𝜔𝜔 𝜔𝜔 = 2𝜋𝜋𝜋𝜋 Ec 1a y 1b.

𝜏𝜏𝑚𝑚 =[1 𝑚𝑚2−1� ]2

𝜔𝜔 Ec. 2.

where f is the modulation frequency of the excitation source. Although for simple systems τm and τφ generally agree, the first is usually shorter than that determined from the modulation factor. In this work, the study of the pH of aqueous samples will be carried out with an optical detection method (phase sensitive measurement) using a luminescent indicator immobilized on a polymeric membrane.


Instruments and materials - Nylon+ membrane doped with the luminescent indicator [Ru(S2D)2 bim]2- (see chemical structure in Fig. 2).

Ru2+

N

N

N

N

N

NHNH

N

-O3S

SO3-

-O3S

-O3S

Figure 2. Chemical structure of the complex [Ru (s2d)2 bim]2-.

- Sodium phosphate monobasic, NaH2PO4. - Sodium phosphate dibasic, Na2HPO4. - Sodium phosphate tribasic, Na3PO4. - pH-meter Crison Meter GLP 22+. - Phase-sensitive luminescence measurements in the laboratory are

carried out with the fiberoptic Optosen multichannel luminescence system. Our Optosen unit uses a high-intensity 470 nm LED as the excitation light source, the output of which is digitally modulated at 90 KHz (user selectable) and is fed through a 430 nm wide band-pass interference filter. The emission from the sensitive terminal is monitored through a 590 nm long-pass filter with a Hamamatsu photodetector module. Synchronous demodulation allows extraction of the emission phase shift. The instrument configuration and data are stored in the Optosen unit but can be transferred at any time to and from a laptop computer via the RS232 or USB ports using the Interlab proprietary software. Remote two-channel temperature data are also collected by the Optosen system to correct the sensor response for temperature effect.

- Pipets of 1.0 and 10.0 mL. - Volumetric flasks of 100 and 250 mL.

Preparation of buffers for the assay (Use Milli-Q purified water or equivalent in all recipes and protocol steps) Preparation of the buffer stock solutions. Prepare three stock solutions

(50 mM) in 250 mL of NaH2PO4, Na2HPO4, Na3PO4, weighting the right amount of the three protonated/deprotonated forms of the sodium phosphates.


Preparation of the buffer work solutions. Using the stocks solutions, mix the right volumes of the stock solutions in order to get 100 mL of 10 mM of phosphate buffer at pH 5, 6, 7, 8, 9.

Protocols Assay protocol. the sensor must conditioned with 80 mL of the sample

during 15-20 min.). During this step the system reaches equilibrium between the sample and the sensor film. This effect can be observed due to a presence of a constant plot after the variation of the signal which depended on the [H+] concentration.

Calibration Curve and sample analysis. Follow the assay protocol for

measuring the phase shift of every work solution and the river water samples.

Calculations and questions

1. Get the calibration curve from the reference solutions used and fit the equation of the curve. Determine the pH of the sample solutions from the calibration problem obtained.

2. Explain the differences between dynamic and static quenching. Justify which type of quenching occurs in the process of pH measurement carried out .

3. Calculate from the data obtained the lifetime of the [Ru (S2d)2 bim]-2

indicator for the following pH measures: 4.0, 5.5, 6.5 , 7.5, 9.0. Justify the luminescent phenomenon that occurs as a function of the lifetimes obtained.

4. Draw the various deprotonated-protonated forms of the complex [Ru (S2D)2 bim]-2 in the studied pH range, indicating the theoretical pKa for each of the reactions taking place .

5. Assuming that the sensor used was working in real time on a river, indicate whether any variation will take place in the sensor measurement from day to night, considering that the pH remains constant during the measurement period. Justify your answer.

References

1. G.Mistlberger, K. Koren, S. M. Borisov, I.Klimant. „Magnetically Remote-Controlled Optical Sensor Spheres for Monitoring Oxygen or pH“. Anal. Chem. 2010, 82, 2124–2128.


2. L.Tormo, N. Bustamante, G. Colmenarejo, G. Orellana. “Can Luminescent Ru(II) Polypyridyl Dyes Measure pH Directly?”. Anal. Chem. 2010, 82, 5195–5204

3. W.J. Bowyer, W. Xu, J.N. Demas. “Determining Proton Diffusion in Polymer Films by Lifetimes of Luminescent Complexes Measured in the Frequency Domain”. Anal. Chem. 2009, 81, 378–384; H.M.R. Goncalves, C.D. Maule, P.A.S. Jorge, Joaquim C.G. Esteves da Silva. “Fiber optic lifetime pH sensing based on ruthenium(II) complexes with dicarboxybipyridine”. Anal. Chim. Acta, 2008, 626, 62–70.

Documents

DEGREE / MASTER chemical... · Optical sensors for oxygen detection ..... 3 4. Optical sensors for pH ..... 5 . 1 . PRACTICAL LESSON 1: Automated portable array biosensor for multisample