Water-Soluble Fluorescent Diblock Nanospheres - American

acrylate)-random-(Texas-red cadaverine succinyloxyethyl methacrylate), PtBA-b-P(HEMA-r-SEMA-r-TX), or poly- mers 1-2 and 2-2: where 1-2 and 2-2 have ...
0 downloads 0 Views 175KB Size
Download PDF
Biomacromolecules 2002, 3, 984-990

984

Water-Soluble Fluorescent Diblock Nanospheres Zhao Li and Guojun Liu* Department of Chemistry, University of Calgary, 2500 University Drive, NW, Calgary, Alberta, Canada T2N 1N4

Say-Jong Law† and Todd Sells† BD Biosciences, 54 Loveton Circle, Sparks, Maryland 21152 Received April 16, 2002; Revised Manuscript Received May 30, 2002

The hydroxyl groups of poly(tert-butyl acrylate)-block-poly(2-hydroxyethyl methacrylate) or PtBA-b-PHEMA were reacted with succinic anhydride to introduce some carboxyl groups into the PHEMA block. Such carboxyl groups were then reacted with Texas-red cadverine (TX-NH2) to incorporate dye molecules. The TX-bearing diblocks formed probably spherical micelles in block-selective solvent DMF/toluene containing 2% DMF. “Permanent” micelles or nanospheres were prepared after cross-linking the TX-bearing PHEMA core block. Such nanospheres were made water soluble by cleaving the tert-butyl groups from the PtBA coronas. Water-soluble nanospheres with high TX numbers and fluorescence quantum yields may find applications in fluorescence in situ hybridization assays. I. Introduction In this paper we report the preparation of water-soluble diblock nanospheres containing many dye molecules in the core. The nanospheres were prepared from two tailor-made diblock copolymers poly(tert-butyl acrylate)-block-poly[(2hydroxyethyl methacrylate)-random-(succinyloxyethyl methacrylate)-random-(Texas-red cadaverine succinyloxyethyl methacrylate), PtBA-b-P(HEMA-r-SEMA-r-TX), or polymers 1-2 and 2-2:

where 1-2 and 2-2 have different n, m, x, and y values as shown in Table 1 and the structure of Texas-red cadaverine (TX-NH2) is

The TX groups were introduced into the diblocks from reacting TX-NH2 with the carboxyl groups of the SEMA units of PtBA-b-P(HEMA-r-SEMA) or Polymers 1-1 and 2-1 (Table 1) via amide bond formation (B f C, Scheme † Currently with Millennium Pharmaceuticals, Inc., 75 Sidney St., Cambridge, MA 02139.

Table 1. Characteristics of the Diblocks Used dnr/dca sample (mL/g) 1-0 1-1 1-2 2-0 2-1 2-2 a

0.095

0.095

h wa LS M (g/mol) 9.5 ×

GPCb NMR M h w/M h n n /m % x % y

104

1.22

1.74

15.9 × 104

1.18

0.98

0 13 9.1 0 18 14.6

0 0 3.9 0 0 3.4

n

m

340 340 340 400 400 400

195 195 195 390 390 390

Measured in CHCl3. b Measured in THF.

1). The PtBA-b-P(HEMA-r-SEMA) diblocks were derived from reacting PtBA-b-PHEMA or polymers 1 and 2 with succinic anhydride (A f B), where 1 had n ) 340 and m ) 195 and 2 had n ) 400 and m ) 390. The PtBA-b-P(HEMAr-SEMA-r-TX) diblocks then formed micelles in DMF/ toluene at a toluene volume fraction of 98% with the P(HEMA-r-SEMA-r-TX) blocks forming the cores (C f D). Nanospheres with PtBA coronas were obtained after crosslinking the TX-bearing PHEMA block with succinyl dichloride. Water-soluble fluorescent nanospheres were obtained after cleaving the tert-butyl groups off the PtBA coronal chains (E f F). Fluorescent polymer micro- or nanospheres have been synthesized traditionally from emulsion1 or dispersion2 polymerization. Dye molecules are loaded into the cores of such spheres by copolymerization1 during or by physical trapping3,4 after sphere preparation. Particles prepared from emulsion polymerization without using polymer surfactant are stabilized by electrostatic repulsion and coagulate at finite salt concentrations.5,6 As will be shown later, this limitation does not apply to the diblock nanospheres prepared here. Although there have been, to our knowledge, no reports on the preparation of fluorescent block copolymer nanospheres, dye tagging has been practiced extensively in the

10.1021/bm025552j CCC: $22.00 © 2002 American Chemical Society Published on Web 07/17/2002

Water-Soluble Fluorescent Nanospheres Scheme 1

past for investigating properties of diblock copolymer micelles. Our group has, for example, labeled the coreforming blocks of diblocks with pyrene to examine the chain exchange kinetics of the micelles.7 Webber and co-workers8 and Winnik and co-workers9 have used dye tagging to investigate various properties of diblock micelles. The preparation of micelles with inherent luminescent blocks has been reported by Jenekhe and Chen.10 The nanospheres reported here are of particular interest because they may be used as probes for applications in fluorescence in situ hybridization (FISH) assays to test for specific DNA or RNA sequences in biological samples.11,12 The nanospheres are water soluble. The surfaces of such spheres bear carboxyl groups that can facilitate the immobilization of various biological polymers including proteins and oligonucleotides. Then, the TX groups absorb in the red end of the visible spectrum, which helps minimize light absorption and emission by inherent chromophores in biological samples and emit fluorescence with high quantum yields. II. Experimental Section Polymer Synthesis and Characterization. Two precursor diblock copolymers, PtBA-b-P(HEMA-TMS), were synthesized by anionic polymerization for this project, where P(HEMA-TMS) denotes poly(2-trimethylsiloxyethyl methacrylate). The procedures used to purify the monomers and to perform the polymerization have been detailed before13 and are thus not repeated here. The diblocks were then hydrolyzed in THF/methanol to yield PtBA-b-PHEMA, which are denoted as polymers 1 and 2. For sample

Biomacromolecules, Vol. 3, No. 5, 2002 985

characterization, the PtBA-b-PHEMA samples were made soluble in common organic solvent such as THF and chloroform by reacting with cinnamoyl chloride to yield PtBA-b-PCEMA, where PCEMA denotes poly(2-cinnamoyloxyethyl methacrylate). The PHEMA block was converted into a PCEMA block, as the esterification step had been utilized in our laboratory extensively in the past and the esterification efficiency was quantitative within experimental error.14 The PtBA-b-PCEMA diblocks, 1-0 and 2-0, were characterized by gel permeation chromatograpy, light scattering, and NMR following procedures detailed before13,14 with results shown in Table 1. PHEMA Labeling with Succinic Groups. To label the PHEMA blocks with succinic groups, the PtBA-b-PHEMA samples were reacted with limiting amounts of succinic anhydride in pyridine at room temperature. To prepare PtBAb-P(HEMA-r-SEMA) or polymer 1-1 with properties shown in Table 1, 0.454 g of polymer, containing 1.29 mmol of hydroxyl groups, was dissolved in 5 mL of pyridine freshly distilled over CaH2. Succinic anhydride, 0.0564 g or 0.564 mmol, was then added. The mixture was stirred overnight before it was dropped over fresh ice crystals to precipitate out the polymer. After pyridine was evaporated off, the sample was filtered and dried to yield a white solid, 0.450 g. Polymer 2-1 was prepared similarly. The succinyl molar fractions x in the PHEMA blocks were obtained from NMR analyses performed in deuterated DMF to be 13% for 1-1 and 18% for 2-1. Texas Red Attachment. Texas Red cadaverine was attached to the succinyl carboxyl groups via amide bond formation catalyzed by 1-(3-dimethylaminopropyl)-3-ethylcarbodiimide hydrochloride (EDCI) following methods reported for coupling other amine and carboxyl groups.15 To label polymer 1-1, 26.0 mg of the sample containing 9.1 × 10-6 mol of succinic groups was dissolved in 0.6 mL of DMF. To the solution was then added half of a mixture consisting of 0.03 mL of 0.10 M Na2HPO3 buffer at pH 7.4, 0.08 mL of water, and 0.14 mL of DMF. The other half of the buffer mixture was used to dissolve TX-NH2, 5 mg or 7.2 × 10-6 mol. EDCI, 8 mg or 0.042 mmol, N-hydroxysulfosuccinimide, 8 mg or 0.037 mmol, and TX-NH2 were then mixed with the polymer solution, and the resultant mixture was stirred at room temperature overnight. The final mixture was dialyzed in a cellulose membrane tube (Molecular Porous) with a molar mass cutoff between 12 000 and 14 000 g/mol against water for 2 days and against distilled THF for 0.5 h to remove low-molar-mass impurities. After most of the solvent was blown off by argon, the sample was dried under vacuum to yield a dark purple solid, 1-2, weighing 27.8 mg. Micelle Formation and Cross-Linking. Micelles of each diblock with PtBA coronas were prepared in DMF/toluene containing 98 vol % of toluene. The HEMA units of the micelles were cross-linked at 80 °C with succinyl chloride. To prepare micelles, 27.8 mg of 1-2 containing ∼0.050 mmol hydroxyl groups was dissolved in 0.38 mL of DMF. Under stirring, 28 mL of toluene was then added gradually. After degassing with Ar, the micellar solution was heated to 80 °C. Triethylamine, 3.7 mg or 0.037 mmol, and succinyl

986

Biomacromolecules, Vol. 3, No. 5, 2002

chloride, 3.7 mg or 0.024 mmol, were added. After 2.5 h at 80 °C, the solution was cooled and the solvents were rotaevaporated to dryness. After extended vacuum-drying, 25.3 mg of solid was obtained. Cleavage of the t-Butyl Groups. The tert-butyl groups of the PtBA coronal chains of the nanospheres were cleaved by treatment with CF3COOH in dry CH2Cl2.16,17 To cleave the tert-butyl groups of 1-2 nanospheres, 6.0 mg of the sample was dissolved in a mixture consisting of 30 mL of CH2Cl2 and 3 mL of CF3COOH. The nanospheres settled after 4 h due to the insolubility of the formed PAA coronal chains. The supernatant was then decanted, and the nanospheres were dissolved in 6 mL of CH3OH/H2O (v/v ) 1/1) containing 0.1 g of Na2CO3. The sample was purified after dialyzing against distilled water for 2-3 days. Purple solid, 4.0 mg, of the sample was obtained after rotaevaporating water and drying under vacuum. DLS and TEM Studies of the Nanospheres. Nanosphere solutions at ∼0.1 mg/mL were used for dynamic light scattering studies (DLS) at a scattering angle of 90°. The DLS of cross-linked micelles was performed in CH2Cl2 and that of the water-soluble nanospheres in a 0.48 M aqueous NaHCO3 solution. The samples were clarified by passing them through 0.2-µm filters. The instrument used (Brookhaven model 9025) was equipped with a He-Ne laser operated at 632.8 nm. The light scattering data were treated following the method of cumulants18 to yield both the hydrodynamic diameter, Dh, and the polydispersity, K2/K12, for the nanospheres. A Hitachi-7000 instrument was operated at 75 kV to obtain the transmission electron microscopy images of the watersoluble nanospheres. Methanol solutions of the spheres were sprayed on to carbon-coated copper grids. A drop of a saturated uranyl acetate solution in ethanol/water (v/v ) 10/ 90) was then dispensed on such a grid to allow the staining of the carboxyl groups of the nanospheres for 15 min before the grid was rinsed three times with distilled water droplets. The staining and rinsing droplets were removed using a piece of facial tissue from the side of the grid opposite to that on which they were dispensed. Fluorescence Measurements. For fluorescence measurements, the excitation and emission slit widths were set at 0.5 and 1.0 nm, respectively. The fluorescent intensities of the TX-labeled chains and cross-linked micelles were compared with those of Texas Red cadaverine at approximately equal absorbances of ∼0.15 in methanol to obtain the relative fluorescence quantum yields φ/φ0. The φ/φ0 values were determined for the water-soluble nanospheres by comparing fluorescence intensities in PH 7.0 buffer containing Na2HPO4 (1.3 × 10-4 M) and KH2PO4 (1.6 × 10-4 M). The φ/φ0 values were reported as averages from duplicate runs. FTIR and NMR Analyses. 1H NMR analyses were performed with a Bruker ACT200 instrument using DMFd7 as solvent. FTIR analyses was performed on pressed polymer-containing KBr disks using a Nexus 470 FTIR instrument.

Li et al.

Figure 1. 1H NMR spectra of polymer 1 in DMF-d7 at different reaction stages: (a) in the PtBA-b-PHEMA form, (b) in the PtBA-bP(HEMA-r-SEMA) or 1-1 form, (c) in the PtBA-b-P(HEMA-r-SEMA) form after TX labeling or as 1-2, and (d) as cross-linked micelles.

III. Results and Discussion PtBA-b-PHEMA Characterization. Two PtBA-b-PHEMA diblocks were synthesized for this project. Shown in Figure 1 is a 1H NMR spectrum of polymer 1. Also shown in the figure are the assignments of the different peaks. From the relative intensities of peaks of different blocks, we obtained n/m ) 1.75/1.00. The other diblock had n/m ) 0.98/ 1.00. For characterizing the samples by gel permeation chromatography (GPC) and light scattering (LS), the diblocks were first reacted with cinnamoyl chloride to yield PtBAb-PCEMA or 1-0 and 2-0. The variation in refractive index differences, ∆nr, between PtBA-b-PCEMA solutions and chloroform was plotted in the form of ∆nr/c vs concentration c for each sample.19 Extrapolating the ∆nr/c data to zero concentration yielded, within experimental error, the same refractive index increment dnr/dc of 0.095 mL/g. (Table 1) for the two samples. The weight-average molar masses of the PtBA-b-PCEMA were determined by light scattering using the Zimm method, and the results are shown in Table 1. The molar masses were apparent, because the dnr/dc values for the two blocks were different and each block had a molar mass distribution.19 While we do not know how to correct for the sample heterogeneity effect to obtain the true molar masses, the correction factor should be approximately 1 as the dnr/dc values for PCEMA and PtBA should be close in CHCl3. Combining the light scattering and NMR results, we obtained the weight-average n and m values of 340 and 195 for polymer 1-0 and 400 and 390 for polymer 2-0, respectively. Attachment of Succinic Groups. After succinic group attachment to the PHEMA block, we obtained 1-1 and 2-1. Shown as Figure 1b is a 1H NMR spectrum of 1-1. The

Water-Soluble Fluorescent Nanospheres

Biomacromolecules, Vol. 3, No. 5, 2002 987

Figure 3. Comparison of visible absorption spectra of (a) TX-NH2 in CH3OH (maximum at 588 nm), (b) 1-2 in CH3OH (maximum at 586 nm), (c) cross-linked micelles of 1-2 in CH3OH (maximum at 588 nm), and (d) water-soluble nanospheres of 1-2 in pH ) 7 buffer (maximum at 590 nm).

Figure 2. FTIR absorption spectra of the polymer 1 series at different reaction stages: (a) in the PtBA-b-PHEMA form, (b) in the PtBA-bP(HEMA-r-SEMA) or 1-1 form, (c) in the PtBA-b-P(HEMA-r-SEMA) form after TX labeling or as 1-2, (d) as cross-linked micelles, and (e) as water-soluble nanospheres.

anticipated reaction between the PHEMA block and succinic anhydride was

The attachment of the succinic groups led to the appearance of two more peaks at 4.2 and 4.4 ppm due to shift in the positions of the pendant oxyethylene protons (or C and D protons of Figure 1). A new peak at 2.6 ppm accounted for the methylene protons of the succinic groups. A more quantitative analysis of the NMR spectra yielded a SEMA molar fraction x of 13% in the PHEMA block for 1-1, which is ∼33% of the intended labeling density. Similar reaction efficiencies were found for other polymers prepared but not described here. The incomplete reaction may be due to the insufficient reaction time and mild reaction conditions used or due to the presence of a trace amount of water not removed from the system. We have also performed FTIR analysis of polymer 1-1 with its IR absorption spectrum shown in Figure 2b for comparison with that of the PtBA-b-PHEMA precursor. The attachment of the succinic groups does not seem to introduce profound changes to the spectrum of the PtBA-b-PHEMA precursor except that carboxyl absorption at 3260 cm-1 became more pronounced. TX Labeling. TX-NH2 was attached to the succinyl carboxyl groups via amide bond formation. The resultant mixture was then dialyzed against distilled water until there was no further leaking out of the dark purple dye or derivatives of TX-NH2 from the dialysis tube. Since the cutoff molar mass for the dialysis membrane was between

12 000 and 14 000 g/mol, the retention of the dye color inside the tube after extended dialysis suggested the covalent attachment of the residual TX-NH2 groups on to the polymer chains. Illustrated in Figure 1c is a 1H NMR spectrum of 1-2. Due to the low content of TX in this sample, the elucidation of bonding between TX and the diblock was difficult. The presence of TX in such a sample was, however, evident from the ripples seen between 6 and 9 ppm of the spectrum at a higher magnification. The signals showed that most of the peaks in this region matched those of TX-NH2. Shown in Figure 2c is a FTIR spectrum of 1-2. The emergence of a new peak at 1652 cm-1, characteristic of amide groups, suggested TX attachment to the diblock by covalent bonds. Shown in Figure 3 is the comparison between the visible absorption spectra of TX-NH2 and 1-2. There is little change in the TX spectrum after attachment to the diblock in the region examined. Assuming that the extinction coefficient of the TX groups at its maximum of 588 nm is the same as that of TX-NH2, we obtained TX labeling molar fractions of 3.9% and 3.4% for 1-2 and 2-2, respectively. These values corresponded to the TX-NH2 utilization efficiencies of ∼36% as shown in Table 2. Compared in Figure 4 are the fluorescence spectra of TXNH2 and 1-2. The maximum of the 1-2 spectrum was blueshifted by 3.0-602 nm. Comparing the peak heights of fluorescence of TX-NH2, 1-2, and 2-2 at a comparable absorbance of ∼0.15 yielded relative fluorescence quantum yields, φ/φ0, of 0.38 ( 0.04 for 1-2 and 0.17 ( 0.03 for 2-2. Micelle Formation and Cross-Linking. PHEMA is not soluble in toluene. After the block is tagged with polar succinyl and TX groups, the PHEMA blocks should be even less soluble in toluene. PtBA, on the other hand, was readily dissolved in toluene. The TX-labeled diblocks should thus form micelles in DMF/toluene with cores consisting of the TX-labeled PHEMA block. Due to the deep color of TX, we were unfortunately not able to see any bluish tint from such solutions associated with micelle formation. We could not obtain TEM images for the micelles either at this stage due to lack of an easy and simple technique to stain the PtBA or PHEMA chains.

988

Biomacromolecules, Vol. 3, No. 5, 2002

Li et al.

Table 2. Relative Fluorescence Efficiencies φ/φ0 of the Diblocks at Different Stages PtBA-b-P(HEMA-r-SEMA-r-TX) sample 1-2 2-2

%

ythea

10.0 10.0

cross-linked micelles

water-soluble spheres

%y

φ/φ0

% y′

φ/φ0

% y′′

φ/φ0

3.9 3.4

0.38 ( 0.04 0.17 ( 0.03

1.9 1.3

0.10 ( 0.01 0.16 ( 0.01

1.2 0.92

0.73 ( 0.03 0.59 ( 0.02

a The variables y, y′, and y′′ denote TX labeling densities measured at different stages of nanosphere preparation. The term y the is the density obtainable if all the TX-NH2 molecules reacted with polymer 1-1 or 2-1.

Figure 4. Comparison of fluorescence spectra of (a) TX-NH2 in CH3OH (maximum at 605 nm), (b) 1-2 in CH3OH (maximum at 602 nm), (c) cross-linked micelles of 1-2 in CH3OH (maximum at 605 nm), and (d) water-soluble nanospheres of 1-2 in pH ) 7 buffer (maximum at 605 nm). The absorbances were not adjusted for different samples.

Assuming micelle formation, we proceeded to their crosslinking using succinyl chloride (Scheme 1). A cross-linking reaction should involve ester bond formation between each succinyl group and hydroxyl groups of two different chains as shown in Scheme 1. A qualitative test for successful crosslinking was to examine the solubility of the resultant micelles in CH2Cl2. Micelles unsuccessfully cross-linked were not dispersed in CH2Cl2 due to the insolubility of the PHEMA block. Using this test, we were able to establish that the minimal amount of time required to cross-link the micelles at 80 °C was ∼2 h and we normally used 2.5 h for this purpose. The need to use a high temperature for PHEMA crosslinking was a surprise to us initially. This is, however, reasonable considering that the glass transition temperature of PHEMA is anywhere between 311 and 393 K depending on its tacticity20 and reagent transport and molecular motion are facilitated only above the glass transition temperature. Shown in Figure 1d is a 1H NMR spectrum of these crosslinked micelles. The PHEMA and TX peaks are now invisible, despite the fact that un-cross-linked TX-bearing PHEMA chains dissolved in DMF-d7. This suggested the low mobility of the TX-bearing chains and supported the hypothesis of insoluble, cross-linked cores. Unfortunately, we were not able to verify PHEMA block cross-linking from FTIR results shown in Figure 2d, as the spectra of the crosslinked and un-cross-linked diblocks were similar. Nanosphere preparation was also supported by our dynamic light scattering results. The hydrodynamic diameters, Dh, of cross-linked 1-2 and 2-2 micelles measured in CH2Cl2 were 81 and 92 nm, respectively. These values are much larger than those of isolated chains. We could not determine the weight-average molar masses of the cross-linked micelles, as the samples absorbed light at the laser wavelength at sufficiently high concentrations. Optical Properties of Nanospheres with PtBA Coronas. One of the more challenging aspects of this project was to

maintain the integrity of the TX groups while ensuring core cross-linking. Compared in Figure 3 was the visible absorption spectrum of 1-2 nanospheres with those of TX-NH2 and 1-2 chains. The TX absorption peak shape remained unchanged after micelle core cross-linking. The disappointing aspect was that the TX contents, as determined from absorption analysis assuming equal molar extinction coefficients for TX in the cores and TX-NH2, decreased by 5060% after micelle core cross-linking (Table 2). This TX content decrease could also be judged by the naked eye, as the color of a sample faded somewhat after core crosslinking. The TX content decrease had to be due to side reactions between succinyl chloride or some reaction products and the TX groups. The TX damage can, in principle, be avoided or reduced in the future by loading the TX groups after nanosphere preparation. We did not attempt this approach as we thought that the TX groups would be more uniformly distributed inside the cores if they were incorporated into the diblock before micelle formation. Even more disappointing was the substantial decrease in the relative fluorescence quantum yields (Table 2) of the residual TX groups in the nanosphere cores. We do not know the exact cause for the fluorescence quantum yield decrease. This decrease was of no practical consequence, however, as it was later recovered after tert-butyl group removal from the PtBA coronal chains and neutralization of the acid used to cleave the tert-butyl groups. Cleavage of t-Butyl Groups. The tert-butyl groups were hydrolyzed from PtBA using trifluoroacetic acid in dry CH2Cl2. Shown in Figure 2e is an FTIR spectrum of the watersoluble nanospheres of 1-2. The disappearance of the rocking band at 845 cm-1 suggests the removal of the tert-butyl groups. This is also demonstrated by the solubility of the resultant nanospheres in water for pH > 5. Properties of the Water-Soluble Nanospheres. Shown in Figure 5 is a transmission electron microscopy (TEM) image of the 2-2 nanospheres. The projections of the particles look circular, suggesting the spherical shape of the particles. The particles look darker on the periphery, because only the PAA chains were stained by uranyl acetate. Averaging the size of more than 50 particles, we obtained an average TEM diameter, DE, of 28.1 ( 1.1 nm for the spheres. Similar TEM images were obtained for the 1-2 nanospheres and the average TEM diameter was 19.6 ( 1.7 nm. We also obtained the hydrodynamic diameters Dh of the spheres in 0.48 M NaHCO3 solution. The Dh values are typically four to five times larger than DE values. This can be partially explained by the swelling of the coronal PAA chains in basic media. The large polydispersity indices, K2/ K12, suggest the aggregation of some nanospheres as well.

Biomacromolecules, Vol. 3, No. 5, 2002 989

Water-Soluble Fluorescent Nanospheres

Table 3. Hydrodynamic and Electron Microscopy Diameters, Dh and DE, and Polydispersity, K2/K12, of the Nanospheres at Different Stages cross-linked micellesa

water-soluble nanospheres

sample

Dh/nm

K2/K12

Dh/nm

K2/K12

DE/nm

1-2 2-2

81 92

0.074 0.192

94 118

0.25 0.29

19.6 ( 1.7 28.1 ( 1.1

a Numbers determined for another set of samples with slightly different TX labeling densities.

Table 4. Effect of Salt Concentration Variation on the Relative Fluorescence Quantum Yields of the Water-soluble 1-2 Nanospheres

Figure 5. TEM image of water-soluble 2-2 nanospheres.

Shown in Figure 4d is a visible absorption spectrum of the 1-2 nanospheres measured in pH 7.0 buffer. The spectrum again does not appear much different from that of TX-NH2. Assuming an equal molar extinction coefficient for the TX groups in the nanospheres and TX-NH2, we obtained the y values of 1.2% and 0.92% for the 1-2 and 2-2 spheres. These numbers represent a ∼30% decrease from those of the nanospheres with PtBA coronas. This decrease either can be from the destruction of more TX groups during tert-butyl group hydrolysis or can be accounted for by molar extinction coefficient variation with solvation medium. Using the TEM diameters of 19.6 and 28.2 nm for the 1-2 and 2-2 spheres and assuming a density of ∼1.0 g/mL for the nanospheres, we obtained 47 and 89 chains for each 1-2 and 2-2 sphere, respectively. These numbers corresponded to 110 and 320 TX groups per sphere. For the spheres to function as reporters in FISH assays, the fluorescence quantum yields should be high. The fluorescence quantum yields of the TX groups in the 1-2 and 2-2 spheres relative to those of TX-NH2, φ/φ0, in pH 7.0 water were 0.73 ( 0.03 and 0.59 ( 0.02, respectively. The fluorescence quantum yields should be high in the cores of the nanospheres, because fluorescence quenching by oxygen and due to thermal motions should decrease relative to the case in water or methanol. The relative quantum yields are not higher than unity as the core may contain quenching groups such as carboxyl groups etc. Furthermore, selfquenching can occur inside the core due to the high local concentration of TX. One of the potential advantages of using block copolymer nanospheres in FISH assays is the stability of such particles against salt-induced aggregation. The current nanospheres with poly(sodium acrylate) coronal chains are stabilized not only by the electrostatic21 but also by the steric stabilization22 mechanism. Shown in Table 4 is the variation of the relative fluorescence quantum yields of 1-2 nanospheres dispersed in water as a function of NaHCO3 concentration. The fact that the spheres remained dispersed in 1.43 M NaHCO3 provides sharp contrast to the behavior of latexes produced from emulsion polymerization without using a polymer surfactant.5,6

NaHCO3 concn (M)

φ/φ0

NaHCO3 concn (M)

φ/φ0

0 0.33 0.64

0.75 0.56 0.53

0.95 1.43

0.53 0.49

Conclusions Two PtBA-b-PHEMA diblocks were prepared. Each diblock was reacted with succinic anhydride to obtain PtBAb-P(HEMA-r-SEMA) diblocks. These were subsequently labeled with TX-NH2 to yield polymers 1-2 and 2-2. Such diblocks containing TX groups formed spherical micelles in DMF/toluene containing 2% DMF. Nanospheres were obtained after cross-linking the hydroxyl groups of the TXlabeled PHEMA core blocks with succinyl chloride. The nanospheres were made water soluble from cleaving the tertbutyl groups off the PtBA coronas. Such nanospheres were shown to be uniform in size by TEM. Even at hundreds of TX groups per nanosphere, the TX fluorescence quantum yields remained high. When attached to oligonucleotide sequences, such nanospheres may function as reporters in FISH assays. Acknowledgment. BD Biosciences is gratefully acknowledged for sponsoring this research. Dr. Xiaohu Yan is thanked for creating Scheme 1. References and Notes (1) See, for example: Dreyer, W. J.; Rembaum, A. In Encyclopedia of Polymer Science and Engineeering, 2nd Ed.; Mark, H. F., Bikales, N. W., Overberger, C. G., Eds.; Wiley & Sons: New York, 1985. (2) See, for example: (a) Margel, S.; Beitler, U.; Ofarim, M. J. Cell Sci. 1982, 56, 157. (b) Melamed, O.; Margel, S. J. Colloid Interface Sci. 2001, 241, 357. (3) Haugland, R. P. Handbook of Fluorescent Probes and Research Chemicals, 10th ed.; Molecular Probes: Eugene, OR, 1996. (4) Eckert, W.; Fuchs, O.; Raizner, F.; Rentel, H. US Patent No. 2,994,697, 1961. (b) Wilson, R. C.; Cranford, N. J.; Freyermuth, H. B. US Patent No. 3,096,333, 1963. (c) Brinkley, J. M.; Haugland, R. P.; Singer, V. L. US Patent No 5,326,692, 1994. (5) Molday, R. S.; Dreyer, W. J.; Rembaum, A.; Yen, S. P. S. J. Cell Biol. 1975, 64, 75. (6) Rembaum, A.; Yen, S. P. S.; Cheong, E.; Wallace, S.; Molday, R. S.; Gordon, I. L.; Dreyer, W. J. Macrmolecules 1976, 9, 328. (7) Smith, C. K.; Liu, G. J. Macromolecules 1996, 29, 2060. (b) Underhill, R. S.; Ding, J.; Birss, V. I.; Liu, G. J. Macromolecules 1997, 30, 8298. (8) Eckert, A. R.; Martin, T. J.; Webber, S. E. J. Phys. Chem. A 1997, 101, 1646. (9) Farinha, J. P. S.; Schillen, K.; Winnik, M. A. J. Phys. Chem. B 1999, 103, 2487. (10) Jenekhe, S. A.; Chen. X. L. J. Phys. Chem. B 2000, 104, 6332.

990

Biomacromolecules, Vol. 3, No. 5, 2002

(11) Hames, B. D.; Higgins, S. J. Gene Probes 1 and 2; IRL Press: Oxford, 1995. (b) Kricha, L. J. Nonisotopic DNA Probe Techniques; Academic Press: San Diego, CA, 1992. (b) Forbes, B. A.; Sahm, D. F.; Weissfeld, A. S. Diagnostic Microbiology, 10th ed.; Mosby: St. Luis, 1998. (12) For localization and quantification of mRNA by FISH see, for example: (a) Hougaard, D. M.; Hansen, H.; Larsson, L.-I. Histochem. Cell Biol. 1997, 108, 335. (b) Cloe¨z-Tayarani, I.; Fillion, G. Brain Res. Protoc. 1997, 1, 195. (c) Angerer, L. M.; Angerer, R. C. Methods Cell Biol. 1991, 35, 37. (13) Liu, G.; Ding, J.; Hashimoto, T.; Saijo, K.; Winnik, F. M.; Nigam, S. Chem. Mater. 1999, 11, 2233. (14) Guo, A.; Tao, J.; Liu, G. J. Macromolecules 1996, 29, 2487. (15) Staros, J.; Wright, R. W.; Swingle, D. M. Anal. Biochem. 1986, 156, 220. (16) Bryan, D. B.; Hall, R. F.; Holden, K. G.; Huffman, W. F.; Gleanson, J. G. J. Am. Chem. Soc. 1977, 99, 2353.

Li et al. (17) Greene, T. W.; Wuts, P. G. M. ProtectiVe Groups in Organic Synthesis, 3rd ed.; Wiley & Sons: New York, 1999. (b) Ma, Q. G.; Wooley, K. L. J. Polym. Sci., A: Polym Chem. 2000, 38, 4805. (18) See, for example: Berne, B. J.; Pecora, R. Dynamic Light Scattering with Applications to Chemistry, Biology, and Physics; Dover Publications: Mineola, NY, 1976. (19) Huglin, M. B. Light Scattering from Polymer Solutions; Academic Press: London, 1972. (20) Brandrup, J.; Emmergut, E. H. Polymer Handbook, 3rd ed.; Wiley & Sons: New York, 1989. (21) See, for example: Hunter, R. J. Foundations of Colloid Science; Oxford University Press: Oxford, 1987; Vol. 1. (22) See, for example: Napper, D. H. Polymeric Stabilization of Colloidal Dispersions; Academic Press: London, 1989.

BM025552J