The Effect of Probiotic Lactobacillus Reuteri Prodentis Consumption on Gingival Crevicular Fluid Inflammatory Response and Metagenomic Profiles of Oral Microbiome in Orthodontic Patients.
Phase: NA
Conditions: Healthy Subjects With Fixed Orthodontics
Sponsor: Trisakti University

ELISA protocol: 
1. Coat microtiter plate wells with 100 μl of the antigen/analyte solution, at a concentration of 
between 1-10 μg/ml in coating buffer. Cover the plate and incubate overnight at 4°C. Wash the 
plate 3 times in wash buffer.  
2. Add 150 μl of blocking solution to each well. Incubate for 1 hour at 37°C. Wash 4 times in 
wash buffer.  
3. Add 100 μl of unconjugated detection antibody (appropriately diluted in wash buffer) to each 
well. Incubate for 1 hour at 37°C. Wash 3 times in wash buffer.  
4. Add 100 μl enzyme-conjugated secondary antibody (appropriately diluted in wash buffer) to 
each well. Incubate for 1 hour at 37°C. Wash 3 times in wash buffer.  
5. Add 100 μl of the appropriate substrate solution to each well. Incubate at room temperature 
(and in the dark if required) for 30 minutes, or until desired color change is attained.  
6. Read absorbance values immediately at the appropriate wavelength or add 50 μl of “stop 
solution”. Gently tap plate to ensure thorough mixing. Measure absorbance within 30 minutes  
7. All procedures will be replicated three times. 
 
 

 
16S or 18S Metagenomic Sequencing Library Preparation using iSeq 100 protocol: 
 
1. Amplicon PCR 
 
This step uses PCR to amplify template out of a DNA sample using region of interest-specific primers 
with overhang adapters attached.  
 
Consumables needed: 
 
Items 
Quantity 
Storage 
Microbial Genomic DNA (5 ng/μl in 10 mM Tris pH 8.5) 
2.5 μl per sample 
‐15° to ‐
25°C 
Amplicon PCR Reverse Primer (1 μM) 
5 μl per sample 
‐15° to ‐
25°C 
Amplicon PCR Forward Primer (1 μM) 
5 μl per sample 
‐15° to ‐
25°C 
2x KAPA HiFi HotStart ReadyMix 
12.5 μl per sample 
‐15° to ‐
25°C 
Microseal 'A' film 
  
  
96‐well 0.2 ml PCR plate 
1 plate 
  
[Optional] Bioanalyzer chip (Agilent DNA 1000 kit catalog 
# 5067‐1504) 
  
  
 
Procedures: 
a.  Set up the following reaction of DNA, 2x KAPA HiFi HotStart Ready Mix, and primers: 
Item 
Volume 
Microbial DNA (5 ng/μl) 
2.5 μl 
Amplicon PCR Forward Primer 1 μM 
5 μl 
Amplicon PCR Reverse Primer 1 μM 
5 μl 
2x KAPA HiFi HotStart ReadyMix 
12.5 μl 
TOTAL 
25 l 
 
b. Seal plate and perform PCR in a thermal cycler using the following program: 
1) 95oC for 3 minutes 
2) 25 cycles of: 
i. 95oC for 30 seconds 
ii. 55oC for 30 seconds 
iii. 72oC for 30 seconds 
3) 72oC for 5 minutes 
4) Hold at 4oC 
 
c. Run 1 μl of the PCR product on a Bioanalyzer DNA 1000 chip to verify the size. Using the V3 
and V4 primer pairs in the protocol, the expected size on a Bioanalyzer trace after the Amplicon 
PCR step is ~550 bp. Other option, using gel electrophoresis to verify the size is also viable. 
Mix 2 μl of the PCR product with 3 μl Blue Juice. Run the sample in 1% agarose gel in 1x TAE 
Buffer with the configuration of 110 V, 15 minutes. With 1kb DNA ladder as the control, the 
band should be produced near ~550 bp. 
 
 
 

2. PCR Clean Up I 
 
This step uses AMPure XP beads to purify the 16S V3 and V4 amplicon away from free primers and 
primer dimer species. 
 
Consumables needed: 
Items 
Quantity 
Storage 
10 mM Tris pH 8.5 
52.5 μl per sample 
‐15° to ‐25°C 
AMPure XP beads 
20 μl per sample 
2° to 8°C 
Freshly Prepared 80% Ethanol (EtOH) 
400 μl per sample 
  
96‐well 0.2 ml PCR plate 
1 plate 
  
[Optional] Microseal 'B' film 
  
  
[Optional] 96‐well MIDI plate 
1 
plate 
  
 
Preparation: 
- 
Bring AMPure XP beads to room temperature 
 
Procedure: 
a. Centrifuge the Amplicon PCR plate at 1,000 × g at 20°C for 1 minute to collect condensation, 
carefully remove seal. 
b. [Optional - for use with shaker for mixing] Using a multichannel pipette set to 25 μl, transfer 
the entire Amplicon PCR product from the PCR plate to the MIDI plate. Change tips between 
samples. 
NOTE 
Transfer the sample to a 96‐well MIDI plate if planning to use a shaker for mixing. If 
mixing by pipette, the sample can remain in the 96‐well PCR plate. 
c. Vortex the AMPure XP beads for 30 seconds to make sure that the beads are evenly dispersed. 
Add an appropriate volume of beads to a trough depending on the number of samples processing. 
d. Using a multichannel pipette, add 20 μl of AMPure XP beads to each well of the Amplicon 
PCR plate. Change tips between columns. 
e. Gently pipette entire volume up and down 10 times if using a 96‐well PCR plate or seal plate 
and shake at 1800 rpm for 2 minutes if using a MIDI plate. 
f. Incubate at room temperature without shaking for 5 minutes. 
g. Place the plate on a magnetic stand for 2 minutes or until the supernatant has cleared. 
h. With the Amplicon PCR plate on the magnetic stand, use a multichannel pipette to remove and 
discard the supernatant. Change tips between samples. 
i. 
With the Amplicon PCR plate on the magnetic stand, wash the beads with freshly prepared 
80% ethanol as follows: 
1) Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each 
sample well. 
2) Incubate the plate on the magnetic stand for 30 seconds 
3) Carefully remove and discard the supernatant. 
j. 
With the Amplicon PCR plate on the magnetic stand, perform a second ethanol wash as follows: 
1) Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each 
sample well. 
2) Incubate the plate on the magnetic stand for 30 seconds. 
3) Carefully remove and discard the supernatant. 
4) Use a P20 multichannel pipette with fine pipette tips to remove excess ethanol. 
k. With the Amplicon PCR plate still on the magnetic stand, allow the beads to air‐dry for 10 
minutes. 

l. 
Remove the Amplicon PCR plate from the magnetic stand. Using a multichannel pipette, add 
52.5 μl of 10 mM Tris pH 8.5 to each well of the Amplicon PCR plate. 
m. Gently pipette mix up and down 10 times, changing tips after each column (or seal plate and 
shake at 1800 rpm for 2 minutes). Make sure that beads are fully resuspended. 
n. Incubate at room temperature for 2 minutes. 
o. Place the plate on the magnetic stand for 2 minutes or until the supernatant has cleared. 
p. Using a multichannel pipette, carefully transfer 50 μl of the supernatant from the Amplicon 
PCR plate to a new 96‐well PCR plate. Change tips between samples to avoid cross‐
contamination. 
 
 
3. Index PCR 
 
This step attaches dual indices and Illumina sequencing adapters using the Nextera UD Indexes Kit. 
 
Consumables Needed: 
Items 
Quantity 
Storage 
2x KAPA HiFi HotStart ReadyMix 
25 μl per sample 
‐15° to ‐25°C 
Nextera™ DNA UD Indexes Set A 
10 μl per sample 
‐15° to ‐25°C 
PCR Grade Water 
10 μl per sample 
  
Microseal 'A' film 
  
  
96‐well 0.2 ml PCR plate 
1 plate 
  
 
Note: The Nextera™ DNA UD Indexes is already premixed, packaged in 96-well plate, and ready to 
use in single pipetting. Pierce the aluminum foil attached to the 96-well plate with a pipette tips, discard 
after piercing. Pipette the needed amount for reaction with new pipette tips. 
 
Procedure: 
a. Set up the following reaction of DNA, Index Primers, 2x KAPA HiFi HotStart ReadyMix, and 
PCR Grade Water:  
Item 
Volume 
DNA from PCR Clean Up I 
5 μl 
Nextera™ DNA UD Indexes Set A 
10 μl  
2x KAPA HiFi HotStart ReadyMix 
25 μl 
PCR Grade Water 
10 μl  
TOTAL 
50 μl 
 
b. Transfer 5 μl the product of PCR Clean-Up I from each well to a new 96-well plate. The 
remaining 45 μl is not used in the protocol and can be stored for other uses. 
c. Gently resuspend the mixture for 10 seconds to mix. 
d. Cover the plate with Microseal ‘A’. 
e. Centrifuge the plate at 1000 x g at 20oC for 1 minute. 
f. Perform PCR on thermal cycler using the following program: 
1) 95oC for 3 minutes 
2) 8 cycles of: 
i. 95oC for 30 seconds 
ii. 55oC for 30 seconds 
iii. 72oC for 30 seconds 
3) 72oC for 5 minutes 
4) Hold at 4oC. 

 
 
4. PCR Clean Up II 
 
This step uses AMPure XP beads to clean up the final library before quantification. 
 
Consumables:  
Items 
Quantity 
Storage 
10 mM Tris pH 8.5 
27.5 μl per sample 
‐15° to ‐25°C 
AMPure XP beads 
56 μl per sample 
2° to 8°C 
Freshly Prepared 80% Ethanol (EtOH) 
400 μl per sample 
  
96‐well 0.2 ml PCR plate 
1 plate 
  
[Optional] Microseal 'B' film 
  
  
[Optional] 96‐well MIDI plate 
1 plate 
  
 
Procedures: 
a. Centrifuge the Index PCR plate at 280 × g at 20°C for 1 minute to collect condensation. 
b. [Optional - for use with shaker for mixing] Using a multichannel pipette set to 50 μl, transfer 
the entire Index PCR product from the PCR plate to the MIDI plate. Change tips between 
samples. 
NOTE 
Transfer the sample to a 96‐well MIDI plate if planning to use a shaker for mixing. If mixing 
by pipette, the sample can remain in the 96‐well PCR plate. 
c. Vortex the AMPure XP beads for 30 seconds to make sure that the beads are evenly dispersed. 
Add an appropriate volume of beads to a trough. 
d. Using a multichannel pipette, add 56 μl of AMPure XP beads to each well of the Index PCR 
plate. 
e. Gently pipette mix up and down 10 times if using a 96‐well PCR plate or seal plate and shake 
at 1800 rpm for 2 minutes if using a MIDI plate. 
f. Incubate at room temperature without shaking for 5 minutes. 
g. Place the plate on a magnetic stand for 2 minutes or until the supernatant has cleared. 
h. With the Index PCR plate on the magnetic stand, use a multichannel pipette to remove and 
discard the supernatant. Change tips between samples. 
i. 
With the Index PCR plate on the magnetic stand, wash the beads with freshly prepared 80% 
ethanol as follows: 
1) Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each 
sample well. 
2) Incubate the plate on the magnetic stand for 30 seconds 
3) Carefully remove and discard the supernatant. 
j. 
With the Index PCR plate on the magnetic stand, perform a second ethanol wash as follows: 
1) Using a multichannel pipette, add 200 μl of freshly prepared 80% ethanol to each 
sample well. 
2) Incubate the plate on the magnetic stand for 30 seconds. 
3) Carefully remove and discard the supernatant. 
4) Use a P20 multichannel pipette with fine pipette tips to remove excess ethanol. 
k. With the Index PCR plate still on the magnetic stand, allow the beads to air‐dry for 10 minutes. 
l. 
Remove the Index PCR plate from the magnetic stand. Using a multichannel pipette, add 27.5 
μl of 10 mM Tris pH 8.5 to each well of the Index PCR plate. 
m. If using a 96‐well PCR plate, gently pipette mix up and down 10 times until beads are fully 
resuspended, changing tips after each column. If using a MIDI plate, seal plate and shake at 
1800 rpm for 2 minutes. 

n. Incubate at room temperature for 2 minutes. 
o. Place the plate on the magnetic stand for 2 minutes or until the supernatant has cleared. 
p. Using a multichannel pipette, carefully transfer 25 μl of the supernatant from the Index PCR 
plate to a new 96‐well PCR plate. Change tips between samples to avoid cross contamination. 
q. (OPTIONAL) Run 1 μl of a 1:50 dilution of the final library on a Bioanalyzer DNA 1000 chip 
to verify the size. Using the V3 and V4 primer pairs in the protocol, the expected size on a 
Bioanalyzer trace of the final library is ~630 bp. Other option, using gel electrophoresis to 
verify the size is also viable. Mix 2 μl of the PCR product with 3 μl Blue Juice. Run the sample 
in 1% agarose gel in 1x TAE Buffer with the configuration of 110 V, 15 minutes. With 1kb 
DNA ladder as the control, the band should be produced near ~630 bp. 
 
5. Library Quantification, Normalization, and Pooling 
 
Illumina recommends quantifying your libraries using a fluorometric quantification method that uses 
dsDNA binding dyes, such as Qubit 3.0. Calculate DNA concentration in nM, based on the size of 
DNA amplicons as determined by Qubit 3.0 Fluorometer using the given formula: 
(𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 𝑖𝑛𝑛𝑔
μl )
(660𝑔
𝑚𝑜𝑙𝑥 𝑎𝑣𝑒𝑟𝑎𝑔𝑒 𝑙𝑖𝑏𝑟𝑎𝑟𝑦 𝑠𝑖𝑧𝑒)
 𝑥 106 = 𝑐𝑜𝑛𝑐𝑒𝑛𝑡𝑟𝑎𝑡𝑖𝑜𝑛 𝑖𝑛 𝑛𝑀. 
For example, in this metagenomic sequencing: 
(15 𝑛𝑔
μl )
(660𝑔
𝑚𝑜𝑙𝑥 630)
 𝑥 106 = 36 𝑛𝑀. 
Dilute concentrated each sample final library using Resuspension Buffer (RSB) or 10 mM Tris pH 8.5 
or Nuclease-Free Water to a concentration 4 nM.  
 
 
6. Final Library Preparation  
 
Preparation:  
a. Thaw the bagged cartridge that was stored in -25oC to -15oC storage. If the cartridge is boxed, 
remove it from the box but do not open the white foil bag. Thaw the bagged cartridge using 
one of the following methods. Use immediately after thawing, without refreezing or otherwise 
storing. 
 
 
Method 
Thaw Time 
Instruction 
20°C to 25°C 
water bath 
6 hours, not 
exceeding 18 
hours 
Use 6 L (1.5 gal) water per cartridge. 
• Set a temperature-controlled water bath to 25°C or mix 
hot and cold water to achieve 20°C to 25°C. 
• Face the bag label up, submerge the cartridge completely, 
and apply ~2 kg (4.5 lbs) weight to prevent floating. 
• Do not stack cartridges in the water bath unless it is 
temperature-controlled. 
2°C to 8°C 
refrigerator 
36 hours, not 
exceeding 72 
hours 
Position the cartridge so that the label faces up and air can 
circulate on all sides, including the bottom. 
Room-
temperature air 
(20°C to 25°C) 
9 hours, not 
exceeding 
18 hours 
Position the cartridge so that the label faces up and air can 
circulate on all sides, including the bottom. 
 

b. While thawing the bagged cartridge, remove the flow cell from 2-8oC storage and set aside the 
unopened package at room temperature for 10-15 minutes to prevent condensation. 
 
Final Library Preparation: 
 
a. Dilute Library to 1 nM in a low-binding tube. Dilute library using Resuspension Buffer (RSB) 
or 10 mM Tris-HCl, pH 8.5 Buffer, or Nuclease-Free Water (NFW).  
b. After dilution, pool all the diluted 1 nM library in one low-binding tube. (OPTIONAL) 
Reconfirm its 1 nM concentration using Qubit 3.0 Fluorometer. 
c. Dilute the pooled 1 nM library to 50 pM concentration (5 μl 1 nM library in 95 μl RSB). Vortex 
briefly, then spin down briefly. Set aside diluted library on ice until sequencing. 
d. Remove 10 nM Phi-X stock from -25oC to -15oC storage. Phi-X is needed only for an optional 
spike-in or a Phi-X only run. 
e.  Thaw Phi-X at room temperature for ten minutes. 
f.  Using RSB/10 mM Tris-HCl pH 8.5 Buffer/NFW, do a two-step dilution of Phi-X from 10 nM 
to 1 nM, and then to 50 pM (5 μl 1 nM Phi-X in 95 μl RSB).  
g.  Combine 19 μl of diluted 50 pM library and 1 μl of diluted 50 pM Phi-X in one tube. Vortex 
briefly and spin down briefly. This mixture is ready for injection to the cartridge. 
 
 
 
7. Loading Consumables into the Cartridge 
 
a.  Open the cartridge bag from the notches. 
b. While avoiding the access window on top of the cartridge, remove the cartridge from the bag. 
Discard the bag. 
c. Invert the cartridge five times to mix reagents. Internal components can rattle during inversion, 
which is normal. 
d. Tap the cartridge (label facing up) on the bench or other hard surface five times to ensure 
reagent aspiration. 
e. Using a new pipette tip, pierce the Library reservoir and push the foil to the edges to enlarge 
the hole, as shown in the picture below. 
 
f. Discard the pipette tip to avoid contamination.  
g. Add 20 μl mixture of diluted library + Phi-X to the bottom of the reservoir. Avoid touching the 
foil. Illustration shown in the picture below.  
            
 

h. Open the flow cell’s white foil package from the notches. Use within 24 hours of opening. 
i. 
Pull the flow cell out of the package. Touch only the plastic when handling the flow cell. Avoid 
touching the electrical interface, CMOS sensor, glass, and gaskets on either side of the glass. 
Illustration is shown in the picture below. 
           
 
j. 
Hold the flow cell by the grip points with the label facing up. 
k. Insert the flow cell into the slot on the front of the cartridge. An audible click indicates that the 
flow cell is in place. When properly loaded, the grip protrudes from the cartridge and the glass 
is visible from the access window. Illustration shown in the picture below. 
              
 
 
A. Loading the flow cell B. Loaded flow cell 
 
l. 
Make sure that the cartridge is thawed and contains the flow cell and diluted library. 
m. Place the cartridge onto the tray so that the access window faces up and the flow cell is inside 
the instrument. Do not push the cartridge or tray into the instrument. Illustration shown in the 
picture below.  
             
 
n. Select Close Door to retract the cartridge and close the door. A panel appears on the left side of 
the screen to show information from the scanned consumables.  
o. Select and edit the run parameters: 
1) To change Read Type, select Single Read or Paired End. 

2) To change Read Cycle, enter 26–151 cycles each for Read 1 and Read 2. Add one cycle 
to the desired number of cycles. 
3) To change the output folder for the current run, enter the path to the location or select 
Browse and navigate to it. 
4) Select Save, which updates the run in both the control software and Local Run Manager. 
p. Select Start Run to initiate the pre-run check. Wait for about 15 minutes for pre-run check to 
complete. The run starts automatically after successful completion. Unless the system is muted, 
a chime sound indicates that the run has started. Monitoring run progress and metrics are 
available as they appear on the Sequencing screen after the cycle of 26.